"Signal Transduction: Protein Phosphorylation". In: Current Protocols ...

CHAPTER 14 

Signal Transduction: Protein 

Phosphorylation 

INTRODUCTION 

Cells use selective phosphorylation of proteins to regulate a vast number of intracellular 

processes. Enzymes termed protein kinases couple phosphate groups to 

tyrosine, serine, or threonine residues in specific amino acid sequence motifs of target 

proteins. The process can be reversed by protein phosphatases. Specificity is determined 

by the ability of each of the many different types of protein kinases and phosphatases to 

recognize specific motifs and proteins. By such site-specific regulation of phosphorylation, 

which affects 10% or more of all proteins, cells can switch on or modulate many 

major signaling and metabolic pathways, as well as regulate cell behavior in biological 

events such as migration and embryonic development. The importance of phosphorylation 

to cell biological regulation is underscored by the fact that cells have over a thousand 

different protein kinases. 

One class of protein phosphorylations regulates enzymes, and the addition or removal of 

a key phosphate activates or suppresses activity of the enzyme. Other protein phosphorylations 

enable a protein to bind to another to form a complex—e.g., via the binding 

of an SH2 domain to a specific tyrosine-phosphorylated site in a target protein. Another 

general cellular strategy is to activate cascades of protein phosphorylation in signaltransduction 

pathways. For example, complex linear and interconnecting pathways of 

sequential phosphorylation of proteins leading to the various types of MAP kinases 

are important regulators of cell growth, differentiation, and gene expression. A current 

overview of this large field of protein phosphorylation is presented in UNIT 14.1, and a 

more specific review of MAP kinase pathways is presented in UNIT 14.3; both provide a 

number of relevant literature references. 

Although phosphorylation has classically been characterized by the incorporation of 32 P 

using radioactive inorganic phosphate, a recent methodological breakthrough of particular 

value to cell biologists involves powerful nonradioactive approaches to the study 

of protein phosphorylation. Studies of complex signaling pathways in cells and tissues 

are now possible even for nonexperts by using immunoblotting and immunofluorescence 

or immunohistochemical methods. These new approaches are based on specific antibodies 

that recognize a phosphate group on one or more amino acids selectively—e.g., 

phosphotyrosine residues on any protein, or the presence of a certain type of phosphate 

linkage on a specific protein such as an activated MAP kinase. A wide selection of these 

immunological tools is now available commercially. UNIT 14.2 provides methods for rapid 

direct characterization of phosphorylated proteins using a specific antibody. If antibodies 

of sufficiently high specificity with respect to a single protein are not available, this unit 

also provides a more indirect approach using immunoprecipitation by antibodies against 

the protein of interest, followed by anti-phosphotyrosine immunodetection. It also describes 

methods for antibody localization of key phosphorylated regulatory molecules. 

This approach permits an investigator to follow the expression patterns or intracellular 

movements of key phosphorylated proteins in cells, or even in various tissues of intact 

organisms. 

Current Protocols in Cell Biology 14.0.1-14.0.3, June 2009 

Published online June 2009 in Wiley Interscience (www.interscience.wiley.com). 

DOI: 10.1002/0471143030.cb1400s43 

Copyright C○ 2009 John Wiley & Sons, Inc. 

Signal 

Transduction: 

Protein 


14.0.1 

Supplement 43

Introduction 

14.0.2 

MAP kinase signaling is central to many critical cell biological regulatory events, and 

UNIT 14.3 provides methods for quantitative characterization of this important signaling 

process. Because of the daunting complexity of signaling via MAP kinases, specific 

antibodies are needed. This unit starts by providing a general protocol for detecting 

MAP kinase activation by immunoblot analysis using specific anti-phospho-MAP kinase 

antibodies. It then provides an alternative approach that uses specific antibodies to isolate 

phosphoprotein, and then phosphorylation of a MAP kinase substrate is measured by 

determining amounts of incorporated 32 P. Because novel kinases may be involved in a 

specific cell biological regulatory event, UNIT 14.3 also presents methods for detecting 

unknown kinases using an in-gel kinase assay. A test substrate is incorporated into an 

SDS-polyacrylamide gel, and electrophoretically separated crude protein extracts are 

evaluated for enzyme bands with ability to produce phosphorylation in vitro. 

Although the recent proliferation of immunological methods for detecting specific types 

of protein phosphorylation has made the analysis of phosphorylation much easier, the 

“gold standard” for characterizing phosphorylation of individual proteins continues to be 

radioactive labeling with 32 P followed by biochemical analysis of the radiolabeled phosphoproteins. 

UNIT 14.4 provides cell culture and biochemical protocols for incorporating 

32 P-labeled inorganic phosphate and for characterizing radiolabeled phosphoproteins. 

UNIT 14.5 presents current methods for unambigously identifying the specific phosporylated 

amino acids in individual phosphoproteins. Appropriate combinations of radioactive 

and immunological approaches should permit full characterization of the protein 

phosphorylation cascades and pathways that regulate many important cell biological 

functions. 

Akt (protein kinase B) is another important kinase, which is regulated by phosphoinositide 

3-kinase downstream of signaling cascades induced by a number of different growth 

factors and integrin ligands. Akt in turn regulates critical cellular functions including cell 

survival, growth, and proliferation, and it can play roles in cancer and other diseases. 

UNIT 14.6 provides protocols for assaying the phosphorylation state of Akt that regulates 

its activity, as well as its dephosphorylation. It also provides a protocol for quantifying 

the translocation of activated Akt to the plasma membrane. 

Cells generally interact with extracellular matrix molecules using integrin receptors, 

which trigger complex signaling cascades that regulate cell survival, growth, migration, 

and differentiation. These cascades involve integrin-associated kinases that include focal 

adhesion kinase (FAK) and c-Src. These and related kinases are both the mediators and 

the targets of specific tyrosine phosphorylation events that play important regulatory 

roles. UNIT 14.7 provides a comprehensive series of protocols for analyzing signal transduction 

involving FAK and the structurally related kinase Pyk2, as well as the master 

regulatory kinase c-Src. Procedures are described for inducing signaling, performing 

immunoprecipitation analysis, measuring kinase activity, and characterizing changes in 

phosphorylation and activity with phospho-specific antibodies. Protocols are also provided 

for visualizing activated FAK, Pyk2, and Src, and there are cell biological assays 

for cell migration regulated by these types of kinase signaling. 

Many cell biological processes are regulated by small GTPases. Cell adhesion, migration, 

and proliferation are strongly regulated by the Rho family of GTPases. Although this 

family consists of a number of members with both distinct and overlapping functions, 

RhoA, Rac, and Cdc42 are particularly important and implicated in a wide range of 

functions. UNIT 14.8 provides methods for determining the activation of each of these key 

Rho GTPases using proteins that bind specifically to the activated form. Specificity of 

these assays is then provided by immunodetection of the specific Rho GTPase isoforms 

bound to these activation-detecting reagents. 

Supplement 43 Current Protocols in Cell Biology

The precise regulation of the activities of small GTPases is important during many biological 

processes. UNIT 14.9 describes detailed protocols for assaying their key regulators, the 

GEFs (guanine nucleotide exchange factors) and the GAPs (GTPase activating proteins). 

These enzyme regulators respectively stimulate or inactivate GTPase activities. Quantification 

of the activities of each of these two types of regulators using these methods can 

provide valuable mechanistic insight into GTPase functions in cell and developmental 

biology. Future units will provide assays for other important cell biological regulators. 

Kenneth M. Yamada 





14.0.3 

Current Protocols in Cell Biology Supplement 43

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The Detection of MAPK Signaling 

The transmission of extracellular signals to their intracellular targets is mediated by a 

network of interacting proteins that relays biochemical messages and thus controls cellular 

processes. Several related intracellular signaling pathways (Seger and Krebs, 1995; 

Chen et al., 2001; Morrison and Davis, 2003; Raman and Cobb, 2003; Rubinfeld and 

Seger, 2004), collectively known as mitogen-activated protein kinase (MAPK) signaling 

cascades, have been demonstrated to play a role in many systems. Transmission of signals 

via these cascades is usually initiated by activation of a small G protein (e.g., Ras) and 

followed by a sequential stimulation of several sets of cytosolic protein kinases. Four 

distinct MAPK cascades (ERK, JNK, p38 and BMK; Fig. 14.3.1) have been elucidated so 

far. Each is named after the subgroup of its MAPK component and is composed of from 

three to five tiers (MAP4K, MAP3K, MAPKK, MAPK and MAPKAPK; Fig.14.3.1), 

where the three tiers MAP3K, MAPKK, and MAPK are considered the core cascade. One 

or more components in each of these tiers phosphorylates and activates components in the 

next level, until a downstream component phosphorylates a target regulatory molecule. 

These cascades can cooperate in transmitting signals from most extracellular stimuli, 

and thus can determine a cell’s fate in response to the ever-changing environment. For 

detailed description see Background Information. 

Because many of the MAPK cascade components are activated by phosphorylation, a 

convenient method to detect their activation is the use of phosphorylation-site-specific 

(anti-phospho) antibodies. Indeed, Basic Protocol 1 describes the use of protein blots 

(immunoblotting; also see UNIT 6.2) to detect protein phosphorylation with anti-phospho 

antibodies (see, e.g., Fig. 14.3.2). This method involves only three steps and usually 

yields impressive results. Unfortunately, the available, reliable anti-phospho antibodies 

are limited to some of the MAPK cascade components, and the use of these antibodies 

does not always reflect the kinetic parameters of the kinase activation. To overcome this 

problem, the unit also includes protocols that utilize the kinase activity of the MAPK 

components to determine their activation (Basic Protocols 2 and 3). Basic Protocol 2 

describes an immunoprecipitation of desired protein kinases followed by phosphorylation 

of specific substrates. Basic Protocol 3 describes an in-gel kinase assay, which is mainly 

used when the identity of the kinase is not known, or when there are no reagents available 

for its determination. 

STRATEGIC PLANNING 

The relative intensity and duration of signals transmitted in each MAPK cascade are 

thought to be major determinants of signaling specificity (Marshall, 1995). Therefore, 

an accurate detection of the amount of signals transmitted via various MAPK cascades 

toward target molecules is important for studying intracellular signaling. Usually, the 

activity of one component of the MAPK level of each cascade (ERK, JNK, p38 and 

BMK) is a sufficient indicator of the transmitted signal. However, sometimes the activity 

of additional components at upstream or downstream levels must be determined because 

of a cross-talk between various cascades. For example, p38 can be activated by as many 

as three distinct MAPKKs (MKK 3, 4, or 6; Ono and Han, 2000), and, therefore, it is 

important to check which one of these MAPKKs is the immediate activator in different 

systems. 

Most components of MAPK cascades belong to the large family of protein kinases, 

which in humans consists of about 520 members (Manning et al., 2002). To study protein 

kinases in general, and MAPK components in particular, specific detection of the activity 

Contributed by Yoav Shaul and Rony Seger 

Current Protocols in Cell Biology (2005) 14.3.1-14.3.34 

Copyright C○ 2005 by John Wiley & Sons, Inc. 

UNIT 14.3 


Transduction 

14.3.1 

Supplement 28

Figure 14.3.1 Schematic representation of MAPK cascades. The ERK cascade is represented in green, the JNK cascade 

in red, the p38 cascade in blue, and the BMK cascade in brown. Components that are shared by more than one cascade 

have combination of colors. The connections between components from different levels are shown by arrows; the specifics 

of these interactions have yet to be defined. This black and white facsimile of the figure is intended only as a placeholder; 

for full-color version of figure go to http://www.interscience.wiley.com/c p/colorfigures.htm. 

The Detection of 

MAPK Signaling 

14.3.2 

of the desired protein kinase is essential. Singling out the activity of a particular protein 

kinase from a multitude of related activities that might mask its activity can be achieved 

in two main ways. One of them requires the use of a specific substrate that is recognized 

only by the desired protein kinase. This method is good for detecting kinases like MEK, 

which seems to specifically and selectively phosphorylate its downstream component, 

ERK. The other, and more common method is to isolate the protein kinase and then use 

a general substrate as an indicator of its activity. This unit will concentrate on the latter 

method of kinase activity determination, which has successfully been used in studies of 

MAPK cascades. 

In one of the first methods used for the systematic detection of protein kinases involved 

in growth factor signaling, protein kinases were isolated using MonoQ fast protein liquid 

chromatography (FPLC; Ahn et al., 1990). This method involves stimulation of tissue 

culture cells, fractionating the cytosolic extracts of these cells on a MonoQ column, and 

examining the resulting fractions for protein kinase activity. Since fractionation with the 

MonoQ column is extremely reproducible, kinases that are activated upon stimulation can 

be detected by comparing the elution profiles of kinases from activated and nonactivated 

cells. The advantages of this method are: (1) the ability to identify novel protein kinases 

and measure their activity, (2) the ability to detect the overall activity of many protein 

kinases, and (3) the method’s good linear range, which allows the determination of 


Figure 14.3.2 Detection of MAPK activity by the methods described in this unit. (A) Detection of 

ERK phosphorylation (activity) with anti–phospho ERK antibody. HeLa cells were grown in 6-cm 

plates until subconfluency and then starved for 18 hr in 2 ml/plate of starvation medium. Two 

plates (indicated) were then pretreated with the MEK inhibitor U0126 (10 µM) for 20 min while 

the other plates were left untreated. The cells were then either stimulated for the indicated times 

with EGF, peroxovanadate (0.1 mM vanadate, 0.2 mM H2O2), or with vehiecle control. Aliquots 

(30 µg) from each sample were separated by a 10% SDS-PAGE and subjected to immunoblotting 

with anti-pERK antibody and anti-gERK antibody (C-16; Santa Cruz Biotechnology), and detected 

using an AP detection system (Sigma). (B) Detection of ERK activity by immunoprecipitation and 

MBP phosphorylation. NIH-3T3 cells were grown in 6-cm plates until subconfluency and then 

starved as described as above. Cells were then stimulated either with VOOH (100 µM sodium 

orthovanadate/200 µM H2O2) for 15 min, with EGF (50 ng/ml) for 5 min, or left untreated (basal). 

Cytosolic extracts were prepared by sonication and the resulting proteins (300 µg) were incubated 

either with 30 µl of protein A beads conjugated with anti-ERK C-terminus antibody (C-16; Santa 

Cruz Biotechnology), represented by (+) or with protein A beads only, represented by (–). The 

phosphorylation reaction on MBP was performed as described in Basic Protocol 2. (C) Detection of 

protein kinase activities by in-gel kinase assay. MCF7 cells overexpressing ErbB-2 receptor were 

stimulated with EGF (50 ng/ml) for the indicated times. The in-gel kinase assay was performed as 

described in Basic Protocol 3. ERK1 and ERK2 bands are indicated. The identity of other bands 

was not determined. 



14.3.3 




14.3.4 

the ratio between the activities of distinct protein kinases at a given time. Although this 

method has been good for some protein kinases, its main disadvantages are that separation 

of various proteins kinases is not always complete, and that it is a very laborious method. 

Another method that is useful in detecting novel protein kinases is the in-gel kinase 

assay (Basic Protocol 3). This technique involves copolymerization of a given substrate 

on a sodium dodecyl sulfate (SDS)–polyacrylamide gel, electrophoresis of the samples 

of interest on the copolymerized gel, and in-gel phosphorylation in the presence of 

[γ- 32 P]ATP. The advantage of this method is that it reveals the molecular weight of 

the kinases with the desired specificity, assisting in the identification of the enzymes of 

interest. Also, several samples can be examined simultaneously. The main disadvantages 

of this procedure are: (1) not all protein kinases can be renatured in the SDS gel, (2) each 

in-gel assay takes 2 or 3 days, and (3) there is a narrow linear range of protein kinase 

activities that can interfere with the detection of the fold induction of protein kinases 

upon stimulation. 

The MonoQ fractionation and in-gel kinase assay methods are mainly used to identify or 

characterize novel protein kinases. However, since the resolutions of these two methods 

are not always adequate and they are very labor-intensive, more specific and convenient 

methods are recommended for the characterization of given protein kinases. Such specific 

methods often require the isolation of the protein kinase of interest, although a 

specific activator or substrate can sometimes be used (as is the case with PKA or MEK). 

The separation is often done by specific antibodies, which can be either directed against 

the phosphorylated sequence (anti-phospho antibodies) or against the whole molecule 

(anti-general antibodies), as described in this unit. The anti-phospho antibody becomes 

useful when the examined protein kinase is activated by phosphorylation and the activating 

phosphorylation site is known. Indeed, this seems to be the case for most MAPKs, 

MAPKAPKs, and MAPKKs known today, but it does not seem to be applicable for 

MAP3Ks and MAP4Ks, as described below. Probably the best known examples are the 

anti–active mitogen-activated protein kinase (MAPK; Seger and Krebs, 1995) antibodies 

(anti-diphospho ERK; Gabay et al., 1997; Yung et al., 1997), which are commercially 

available (Table 14.3.1). In response to stimulation, ERK1 and ERK2 are rapidly phosphorylated 

at up to six sites (Robbins and Cobb, 1992). Phosphorylation of two of these 

sites (threonine 183 and tyrosine 185 in ERK2; Payne et al., 1991) can lead to full activation 

of the enzyme, whereas the effects of phosphorylation of the other sites are not 

yet known. Antibodies directed towards the activation motif of ERK, which is PT-E-PY, 

serve as good tools for detecting its enzymatic activity, whereas anti-PAA antibodies can 

detect the total phosphorylation, which does not fully correlate with ERK’s enzymatic 

activity. An important advantage of using anti-phospho antibodies is that they shorten 

and simplify the procedure of detection, which can be achieved in only one step (immunoblotting 

or cell staining). However, the antibodies do not usually serve as reliable 

tools to determine the exact kinetic parameters of the MAPK, as their dynamic range of 

detection is limited. 

The successful use of sequence-specific anti-phospho antibodies relies on their specificity 

for the phosphorylated form of the examined protein. Monoclonal antibodies, 

which usually confer better specificity than polyclonal ones, are considered a reliable 

tool for distinguishing phosphorylated from nonphosphorylated forms of proteins, although 

affinity-purified polyclonal antibodies can be used as well. Both monoclonal and 

polyclonal antibodies can be generated by immunization with whole phosphorylated 

proteins or with KLH- or BSA-coupled peptides (see UNIT 16.6). Standard procedures for 

the isolation of the monoclonal and polyclonal antibodies are then employed. In this unit, 

the use of anti-phospho antibodies in immunoblot analysis is described. However, most 

of these antibodies can be used also for cell staining, which is described in UNITS 4.3 & 14.2. 


Table 14.3.1 Antibodies Available Against MAPKs 

Tier Name MAPK activated 

Mol. wt. 

(kDa) 

Commercially 

available antibodies 

Supplier a Reference 

MAP4K GCK JNK 91 General Many 

companies 

Pombo et al. (1995) 

GLK JNK 104 General 2, 11 Diener et al. (1997) 

GCKR JNK 100 General 11 Shi and Kehrl (1997) 

HPK JNK 97 General 7 Kiefer et al. (1996) 

HGK JNK 130 General 11 Yao et al. (1999) 

KHS JNK 95 General 11 Tung and Blenis (1997) 

SLK JNK 210 General 2 Sabourin and Rudnicki 

(1999) 

MAP4K4/ 

NIK 

JNK 100 General + phospho Many 

companies 

Su et al. (1997) 

SPAK p38 64 General 2 Johnston et al. (2000) 

MST1 JNK, p38 59 General + phospho 7 Graves et al. (1998) 

TNIK JNK 160 General 2,4 Fu et al. (1999) 

NESK JNK 175 — — Nakano et al. (2000) 

MINK JNK, p38 155 General 2, 8 Dan et al. (2000) 

PAK1 JNK, p38 (?) 68 General + phospho 11 Zhang et al. (1995) 

PAK5 JNK 90 General 11 Dan et al. (2002) 

MST4 ERK (?) 52 General 7 Lin et al. (2001) 

MAP3K ASK1 JNK, p38 155 General + phospho Many 

companies 

Ichijo et al. (1997) 

ASK2 JNK 145 General + phospho — Wang et al. (1998) 

TAK1 JNK, p38 82 General + phospho Many 

companies 

Moriguchi et al. (1996) 

LZK1 JNK 140 — — Sakuma et al. (1997) 

DLK1 JNK, p38 110 — — Fan et al. (1996) 

MLK1 JNK 130 General 11 Xu et al. (2001) 

MLK2 JNK, p38-? ERK1/2-? 115 General 11 Hirai et al. (1997) 

MLK3 JNK, p38 ERK-? 93 General + phospho 7, 11 Rana et al. (1996) 

MLK4 JNK, p38 120 — — Gallo and Johnson 

(2002) 

MLTK α/β JNK, p38, ERK1/2-?, 

BMK-? 

95/50 General 14 Gotoh et al. (2001) 

ZAK JNK 91 General 2, 14 Yang (2002). 

MEKK1 JNK, p38. ERK1/2-? 195 General 1, 11 Yan et al. (1994) 

MEKK2 JNK, p38 BMK 70 General 11 Blank et al. (1996) 

MEKK3 JNK, p38, BMK 71 General 11 Blank et al. (1996) 

MEKK4 JNK 180 General 11 Gerwins et al. (1997) 

continued 



14.3.5 


Table 14.3.1 Antibodies Available Against MAPKs, continued 

Tier Name MAPK activated 



14.3.6 

Mol. wt. 

(kDa) 

Commercially 

available antibodies 

Supplier a Reference 

TPL2 JNK, p38, 

ERK1/2, BMK 

60 + General + phospho 7 Salmeron et al. (1996) 

Raf-1 ERK1/2 74 General + phospho Many 

companies 

Kyriakis et al. (1992) 

B-Raf ERK1/2 94 General 1, 11 Peraldi et al. (1995) 

A-Raf ERK1/2 68 General 7 Hagemann and Rapp 

(1999) 

MOS ERK1/2 39 General 1, 11 Posada et al. (1993) 

TAO1 JNK, p38 140 General 4 Hutchison et al. (1998) 

TAO2 JNK, p38 120 General + phospho 2, 9 Chen et al. (1999) 

MAP3K6/7, 

TAO3 

? Not 

charachterized 

— — Manning et al. (2002) 

MAPKK MEK1/2 ERK1/2 45/46 General + phospho Many 

companies 

Ahn et al. (1991) 

MKK3/6 p38 40/41 General + phospho Many 

companies 

Derijard et al. (1995) 

MKK4 JNK, p38 44 General + phospho Many 

companies 

Yan et al. (1994) 

MKK5 BMK 45 General 11 Zhou et al. (1995) 

MKK7 JNK 48 General + phospho Many 

companies 

Tournier et al. (1997) 

MAPK ERK1/2 ERK1/2 42.44 General + phospho 1-14 Ray and Sturgill (1987) 

p38 α−δ p38 41,43.47, 

54 + other 

forms 

General + phospho Many 

companies 

Han et al. (1994) 

JNK1-3 JNK 46,52,54 General + phospho Many 

companies 

Derijard et al. (1994) 

BMK BMK 110 General + phospho Many 

companies 

Zhou et al. (1995) 

MAPKAPKRSK1-3 ERK1/2 90 General + phospho Many 

companies 

Sturgill et al. (1988) 

MAPKAPK2p38 45+other General + phospho Many 

companies 

Stokoe et al. (1992) 

MAPKAPK3p38, ERK1/2 43 General 3, 13 McLaughlin et al. (1996) 

MK5 p38, ERK1/2 56 — — Ni et al. (1998) 

MNK p38, ERK1/2 52 General + phospho 7, 11 Waskiewicz et al. (1997) 

MSK p38, ERK1/2 90 General + phospho 7, 13 Deak et al. (1998) 

SGK BMK 54 General + phospho 11, 13 Hayashi et al. (2001) 

aSuppliers: 1, Abcam (http://www.abcam.com/); 2, Abgent (http://www.abgent.com); 3, Abnova, (http://www.abnova.com/tw); 4, BD Biosciences; 5, 

Biomol; 6, Calbiochem; 7, Cell Signaling (http://www.cellsignal.com); 8, Novus Biologicals; 9, PhosphoSolutions (http://www.phosphosolutions.com); 

10, Promega; 11, Santa Cruz Biotechnology; 12, Sigma; 13, Upstate Biotechnology; 14, Zymed. See SUPPLIERS APPENDIX for additional contact 

information. 


Because not all MAPK components are activated by phosphorylation, and because the 

availability of commercial anti-phospho antibodies might be limited, the activity of 

the desired protein kinases may be determined by immunoprecipitation with specific 

antibodies directed to the C-terminal domain of the kinase (as described in Basic Protocol 

2). Another use for anti-general antibodies is detection of slower migration on SDS-PAGE 

that occurs upon phosphorylation of regulatory residues of some MAPKs. However, this 

gel shift does not always correlate with enzymatic activity, as was shown for ERK and 

Raf1. Methods for affinity purification that do not involve antibodies can sometime be 

used to isolate given protein kinases (e.g., JNKs; Hibi et al., 1993). Although affinity 

techniques (including immunoprecipitation) are often used, it should be noted that the 

attachment to a solid support that occurs in this method may interfere with the accurate 

detection of the kinase activity. However, affinity reagents are not available for all MAPK 

components, and the identity of the desired protein kinase might be obscured. Therefore, 

the method that can be used in this case is the in-gel kinase assay as described in 

Basic Protocol 3. Although other methods are available, the combination of the methods 

described here should allow determination of the activity of most MAPK components 

and identification of the components that are activated by various extracellular stimuli. 

IMMUNODETECTION OF MAPK ACTIVATION USING ANTI-PHOSPHO 

MAPK ANTIBODIES 

This method describes the immunodetection of phosphoproteins using extraction of 

proteins by sonication followed by immunoblot analysis. Although phosphorylation of 

ERK1/2 is used here as an example, this immunodetection protocol can be used with most 

specific anti-phospho antibodies available for many components of the MAPK cascades 

(Table 14.3.1). This protocol minimizes the time phosphorylated proteins are exposed 

to phosphatases, which allows reliable and quantitative detection of the phosphorylated 

proteins, and it can be completed within 7 to 10 hr. Furthermore, this method can be used 

with almost all tissue culture cell lines, homogenized animal organs, and even whole 

lower organisms. 

Materials 

Rat1 cells (ATCC #CRL-2210) 

DMEM containing 10% heat-inactivated FBS (see APPENDIX 2A and UNIT 1.2) 

Starvation medium: DMEM containing 0.1% (v/v) heat-inactivated FBS (see 

APPENDIX 2A and UNIT 1.2) 

50 µg/ml epidermal growth factor (EGF) in EGF buffer (0.5 mg/ml BSA in PBS) 

EGF buffer: 0.5 mg/ml bovine serum albumin (BSA) in phosphate-buffered saline 

(PBS; APPENDIX 2A) 

Phosphate-buffered saline (PBS; APPENDIX 2A), ice-cold 

Buffer A (see recipe), ice-cold 

Buffer H (see recipe), ice-cold 

Protein standards: 5, 10, 20, 50, 100, and 200 µg/ml BSA in Buffer H (see recipe 

for buffer H) 

Bradford protein assay reagent (Pierce or Bio-Rad; also see recipe for Coomassie 

dye reagent in APPENDIX 3H) 

4× sample buffer for SDS-PAGE (see recipe) 

1.5 M Tris·Cl, pH 8.8 (APPENDIX 2A) 

30% acrylamide/0.8% bisacrylamide (Table 6.1.1) 

10% (w/v) ammonium persulfate (prepare fresh) 

Tetramethylethelendiamine (TEMED) 


Running buffer (see recipe) 

Prestained protein markers 

BASIC 

PROTOCOL 1 



14.3.7 




14.3.8 

Transfer buffer (see recipe) 

Blocking solution: 2% (w/v) bovine serum albumin (BSA) in TBST (see recipe for 

TBST) 

Primary antibodies: monoclonal mouse anti-phospho ERK antibody (Table 14.3.1) 

and polyclonal rabbit anti-general ERK antibody (C-terminus) 

Tris-buffered saline with Tween 20 (TBST; see recipe) 

Secondary antibodies: alkaline phosphatase (AP)–coupled goat anti-mouse 

antibody and horseradish peroxidase (HRP)–conjugated goat anti-rabbit 

antibody 

6-cm tissue culture dishes 

1-ml pipet tips, precooled 

1.5-ml microcentrifuge tubes precooled; four sets of six (each labeled 1 to 6) 

Plastic (or rubber) policeman 

Probe sonicator (e.g., Branson) 

96-well flat-bottom microtiter plate 

Microtiter plate reader capable of reading at 595 nm 

95◦C or boiling water bath 

Gel-casting apparatus: 7 × 10–cm glass plates, 1.5-mm spacers, and 1.5-mm comb 

with 10 teeth (also see UNIT 6.1) 

Gel electrophoresis apparatus and power supply (also see UNIT 6.1) 

0.45-µm nitrocellulose membrane (e.g., Schleicher & Schuell or Millipore), cut to 

gel size 

Whatman 3MM paper (two sheets cut to gel size) 

Transfer apparatus and power supply (also see UNIT 6.2) 

Additional reagents and equipment for cell culture (UNIT 1.1), SDS-PAGE (UNIT 6.1), 

and visualization of immunoblotted proteins (UNIT 6.2 & 14.2) 

NOTE: All reagents and equipment coming into contact with living cells must be sterile, 

and aseptic technique should be used accordingly. 

NOTE: All culture incubations should be performed in a humidified 37 ◦ ,5%CO2 incubator 

unless otherwise specified. Some media (e.g., DMEM) require altered levels of 

CO2 to maintain pH 7.4. 

Prepare cellular extracts 

1. Grow Rat1 cells in six 6-cm tissue culture dishes in 4 ml DMEM containing 10% 

FBS to subconfluency (∼0.5 × 10 6 cells/dish). 

UNIT 1.1 describes basic cell culture techniques. 

2. Serum-starve the Rat1 cells by removing the culture medium from each dish, replacing 

it with 2 ml starvation medium, and incubating 18 hr. 

During serum starvation, place the dishes in the tissue culture incubator, making sure that 

the dishes remain flat and that the medium covers the entire dish evenly. The aim of this 

starvation is to make the cells quiescent, and thereby reduce the amount of the inducible 

MAPK phosphatases (MKPs). Under these conditions, this can be achieved within 14 to 

24 hr. Starvation for too long or any change in temperature or pH may be stressful to 

the cells, thereby inducing activation of one or more signaling pathways. Although this 

protocol describes EGF stimulation of Rat1 cells, this procedure, with minor changes, 

can be used for most extracellularly stimulated cells. 

3. To each of three of the dishes, add 2.0 µl of50µg/ml EGF (stimulated samples). 

To each of the other three dishes, add 2.0 µl EGF buffer (control samples). Incubate 

one sample and control for 5 min, another sample and control for 15 min, and the 

third sample and control for 45 min. 


Usually, the stimulus is given first to the dishes with the longest incubation, then, at 

appropriate intervals, to the dishes with the second longest and the shortest incubation 

periods. It is useful to make and use a time chart so that stimuli will be given at the 

appropriate times and the cells harvested within a short period of time (within 5 to 

10 min for all dishes). If the influence of the stimulating agent on the particular cells 

is not yet known, a positive control should be included, such as a dish treated with 

50 µl peroxovanadate (VOOH) solution (see recipe), for a final concentration of 100 µM 

sodium orthovanadate and 200 µMH2O2, which nonspecifically activates many signaling 

events in most tissue culture cells. 

4. At the end of the assay, remove the medium from the dishes containing the Rat1 

cells. Rinse dishes twice with 5 ml ice-cold PBS and once with 5 ml ice-cold buffer 

A. Be sure to remove all of the PBS after the last wash. Place dishes on ice. 

Because the objective at this stage is to arrest or slow down biological processes, the 

dishes should be placed on ice. Washing and harvesting of each dish should take 0.5 to 

1.5 min. so that all six dishes are harvested within 5 to 10 min. 

5. Add 250 µl of ice-cold buffer H to each dish, tilt the dish gently (preferably on 

ice), and scrape the cells into the buffer using a plastic (or rubber) policeman. Using 

precooled pipet tips, transfer the cells and buffer to prelabeled, precooled 1.5-ml 

plastic test tubes. 

Special consideration should be given to the composition of the buffer H (see Reagents 

and Solutions). The authors recommend using β-glycerophosphate, which serves both 

as a buffer and a general phosphatase inhibitor, rather than Tris or HEPES. Sodium 

orthovanadate is used to inhibit tyrosine phosphatases, and the mixture of pepstatin A, 

aprotinin, leupeptin, and benzamidine is used to inhibit proteinases. This buffer, when 

cold, blocks most of the phosphatase and proteinase activities in cell extracts. 

6. Disrupt the cells by sonication using two 7-sec, 50-W pulses with a 20-sec interval 

per ∼0.5-ml sample on ice. 

Over the years, several methods of protein extraction from cells have been successfully 

used in the study of protein kinases. In this protocol, which utilizes sonication, proteins are 

extracted from the cytosolic and nuclear fractions, but not from the membrane fraction, 

and therefore can be considered as cytosolic extract. Cellular extraction with nonionic 

detergents, which extract proteins from membranal, cytosolic, and some nuclear fractions 

of the cell, makes determination of the protein concentration somewhat difficult, but it is 

often used. Extraction with RIPA buffer or by freezing-thawing can also be used for some 

kinases. 

7. Microcentrifuge the cellular extracts 15 min at 14,000 × g, 4 ◦ C. Transfer the resulting 

supernatants (cytosolic extracts), which contain the protein to be examined 

for phosphorylation, to fresh, precooled microcentrifuge tubes. Take a 5- to 20-µl 

aliquot of each extract and determine protein concentration (steps 8 to 11). Store 

the remainder of each cytosolic extract on ice until needed for the electrophoresis, 

transfer, and detection steps (steps 12 to 36). 

The protein concentration of each sample should be determined so that the amounts of 

proteins from the different samples can be compared, making it possible to determine the 

relative amounts of protein kinases in all samples accurately. If samples are compared 

based on cell number, differences of up to 20% in the amount of protein may result. Such 

differences may cause even larger ones in the following steps. 

Determine protein concentration and prepare extracts for electrophoresis 

8. Dilute 10 µl of each cytosolic extract sample in 190 µl buffer A (a final dilution of 

1:20) in labeled tubes. 

Usually, dilutions of at least 1:20 are necessary to ensure that the samples will be in the 

linear range of the protein determination assay. For some Coomassie blue reagents with 

extended ranges, this dilution is not always necessary. 



14.3.9 




14.3.10 

9. Put 10 µl of each of the protein standards (10, 25, 50, 75, 100, 150, and 200 µg/ml 

BSA in buffer H) into duplicate or triplicate wells of a flat-bottom 96-well microtiter 

plate. 

Protein standards should be prepared in the same buffer used for the cell extraction (in 

this case, buffer H). 

10. Put 10 µl of each of the sample dilutions from step 8 into duplicate or triplicate wells 

of the same microtiter plate. Add 200 µl of Bradford protein assay reagent to all 

wells. 

11. Place the microtiter plate in a microtiter plate reader and measure the optical density 

at 595 nM. 

Perform electrophoresis 

12. From the optical densities, calculate the protein concentrations of the samples. Take 

an aliquot of each extract corresponding to 40 µg of protein and place in a new 

1.5-ml microcentrifuge tube. 

30 to 50 µg protein should be loaded in each lane of the gel. The authors strongly 

recommend using equal protein amounts for each sample to avoid inaccuracies. 

13. Add 1/3 vol of 4× sample buffer to each tube, mix the contents, and place in 95 ◦ C 

or boiling water bath for 5 min. 

14. Assemble 7 × 10–cm glass plates and 1.5-mm spacers for casting the polyacrylamide 

gel. 

15. Prepare a 12% separating gel by mixing the following (total volume, 10.0 ml): 

3.4 ml H2O 

2.5ml1.5MTris·Cl, pH 8.8 

4.0 ml 30% acrylamide/0.8% bisacrylamide 

100 µl 10% ammonium persulfate 

6 µl TEMED. 

16. Pour the separating gel (UNIT 6.1). Overlay the top of the gel with water and allow it 

to polymerize ∼30 to 45 min. 

17. Prepare 3% polyacrylamide stacking gel by mixing the following (5 ml total): 

550 µl 30% acrylamide/0.8% bisacrylaimde solution 

625 µl 0.5MTris·Cl, pH 6.8 

3.7 ml H2O 

120 µl ammonium persulfate 

5 µl TEMED. 

18. Remove the water from the top of the polymerized separating gel. Using a Pasteur 

pipet, layer 2 ml of the stacking gel on top of the separating gel (UNIT 6.1) to fill up 

the apparatus. Insert comb into the stacking gel and allow it to polymerize for ∼10 

to 20 min. 

19. Place the polymerized gel in the electrophoresis apparatus, add running buffer, and 

check for leaks. 

20. Load 30 µl of the samples (from step 13) and 10 µl of the prestained protein markers 

into each well of the gel. 

Usually, prestained markers are loaded into the first or second lane of the gel so that the 

first lane can be located in the immunoblot and the molecular weights of the detected proteins 

determined. These markers will also indicate whether the proteins were completely 

transferred from the gel onto the nitrocellulose paper during blotting. 


21. Connect the wire leads of the gel apparatus to the power supply. Turn on the power 

supply and run the gel at 150 V (constant voltage) until the bromphenol blue dye 

reaches a point 0.5 cm from the bottom of the gel (which usually takes ∼1 hr). 

Transfer the proteins to a membrane 

22. Prewet (soak) the nitrocellulose membrane and 3MM Whatman paper in transfer 

buffer. 

23. Once the dye front of the SDS-PAGE has reached the end of the gel, remove the 

gel from the apparatus, cut off the stacking gel, and place the separating gel in a 

container with transfer buffer. 

24. Fill the transfer apparatus with transfer buffer. Open the inner transfer apparatus 

and remove air bubbles from the pads. Make a sandwich by putting a wet 3MM 

Whatman paper on the wet pad, the gel on top of the 3MM Whatman paper, the wet 

nitrocellulose membrane on top of the gel, and the other wet 3MM Whatman paper 

on top of the nitrocellulose membrane. 

Basic immunoblotting techniques are described in detail in UNIT 6.2. 

25. Remove any air bubbles from between the different layers of the transfer sandwich 

by gently rolling a 10-ml pipet over the sandwich. Place the other wet pad on top of 

the transfer sandwich. 

Make sure that no air bubbles are trapped between the gel and the other components. 

26. Place the sandwich containing the gel and nitrocellulose membrane into the bufferfilled 

transfer apparatus with the nitrocellulose membrane facing the side with the 

cathode and the gel facing the side with the anode. Connect the apparatus to a power 

supply and start the current. Transfer the proteins at 200 mA constant current for 

120 min, preferably with a cooling device. 

The voltage will drop as the transfer progresses due to an increase in the conductivity. 

Methanol or 0.05% SDS are sometimes included in the transfer buffer; their inclusion 

will require different transfer conditions (see UNIT 6.2). 

27. At the end of the transfer period, turn off the power supply and remove the nitrocellulose 

membrane from the transfer sandwich. Rinse the nitrocellulose membrane 

with transfer buffer to remove any adhering pieces of gel, and place the membrane 

in a flat container. 

At this stage, the efficiency of protein transfer can be monitored visually by the transfer of 

prestained protein markers to the membrane. The total amount of protein transferred can 

also be detected by staining the membrane with Ponceau S solution (UNIT 6.2). However, 

since the total amount of nonphosphorylated proteins is determined by general antibodies, 

as described later, staining with Ponceau S is probably not essential for this particular 

protocol. 

Expose the membrane to antibodies 

28. Incubate the nitrocellulose membrane in ∼25 ml blocking solution for 60 min at 

room temperature. 

This will ensure that nonspecific protein binding sites on the membrane are blocked. The 

use of milk for blocking is not recommended because it causes nonspecific binding of 

some of the antibodies. 

29. Incubate the blot with 10 to 20 ml of the primary antibody (e.g., monoclonal antiphospho 

ERK antibody, diluted according to the supplier’s recommendations in 

TBST) either overnight at 4 ◦ C, 30 min at 37 ◦ C, or 1 to 2 hr at room temperature. 

30. Wash the blot in the flat container at least three times, each time for 15 min with 50 

ml TBST at room temperature. 



14.3.11 


BASIC 

PROTOCOL 2 



14.3.12 

31. Incubate the blot with 10 to 20 ml of the secondary antibody (e.g., AP-conjugated 

goat anti-mouse IgG, diluted according to the supplier’s instructions in TBST) for 

45 min at room temperature. 

For detection of the anti-phospho ERK antibody, the AP detection system is recommended 

because of its broader linear range compared to ECL (see Commentary). Alternatively, 

HRP-conjugated goat anti-rabbit antibodies can be used as secondary antibodies with 

detection by ECL. If HRP-rather than AP-conjugated antibodies are used at this stage, 

then AP-conjugated antibodies should be used in step 35. 

32. Wash the blot at least three times, each time for 10 min, with 50 ml TBST. 

33. Use an AP detection protocol (UNIT 6.2) to detect phospho ERK. 

After detecting the phosophorylated (active) ERK, it is recommended that it be determined 

whether there is an equal amount of ERK in all lanes by exposing the same blot to 

polyclonal anti-general ERK antibody and redeveloping it (see steps 34 to 36). 

Expose the membrane to general antibody 

34. Incubate the stained nitrocellulose in blocking solution for 30 min at room temperature. 

Since two different types of antibodies are used (mouse and rabbit), stripping away 

the antibodies used in steps 29 to 30 is not necessary, and the second blotting can be 

performed by simply adding the different type of antibody and following steps 35 and 

36. Antibodies from the same species of origin (mouse or rabbit) can be used for both 

steps; however this requires a stripping step, which must be performed before continuing 

with step 35. Note that some of the antibodies can hinder recognition by other antibodies 

directed against different epitopes, and therefore also require stripping before the second 

type of antibody is applied. 

35. Incubate the blot with 10 to 20 ml of the polyclonal rabbit anti-general ERK primary 

antibody, wash as described in step 30, then incubate with 10 to 20 ml of HRPconjugated 

goat anti-rabbit secondary antibody. 

36. Use a luminescent detection protocol for HRP (UNIT 6.2) to observe general ERK. 

The ECL protocol involves mixing solution A (2.5 mM Luminol, 400 mM p-coumarin, 

100 mM Tris·Cl, pH 8.5) with an equal amount of solution B (5.4 mM H2O2, 100 mM 

Tris·Cl, pH 8.5), incubation of the blot with the substrate solution for 1 min, drying 

the blot with 3MM Whatman paper, wrapping the blot in transparent plastic wrap and 

exposing it to X-ray film. See UNITS 6.2 & 14.2 for additional details. 

DETERMINATION OF MAPK (ERK) ACTIVITY BY 

IMMUNOPRECIPITATION 

This method describes determination of ERK activity by isolating the enzyme using 

immunoprecipitation with specific antibodies and then performing a phosphorylation 

reaction in vitro. Although this protocol uses ERK with its appropriate reagents, it can be 

performed with other components of the MAPK cascade, as antibodies are available for 

most components of these cascades (Table 14.3.1). This protocol allows a fast and efficient 

isolation of the desired protein kinase and its reliable quantitation by a phosphorylation 

reaction. It involves standard biochemical procedures and can be completed in 7 to 

10 hr. Furthermore, this method can be used with almost all tissue culture cell lines, 

homogenized animal organs, and even lower organisms. 

CAUTION: When working with radioactive materials, take appropriate precautions to 

avoid contamination of the experimenter and the surroundings. Carry out the experiment 

and dispose of wastes in an appropriately designated area, following guidelines provided 

by the institutional Radiation Safety Officer (also see UNIT 7.1 and APPENDIX 1D). 


Materials 





50 µg/ml epidermal growth factor (EGF) in EGF buffer 

EGF buffer: 0.5 mg/ml bovine serum albumin (BSA) in phosphate buffered saline 





Protein standards: 5, 10, 20, 50, 100 and 200 µg/ml BSA in Buffer H 



Protein A–Sepharose beads (Amersham Biosciences; 10 to 20 µl of packed beads 

per reaction) 

Antibody for immunoprecipitation (e.g., anti-ERK C-terminus antibody, 1 to 5 µg 

per reaction according to the supplier’s instructions) 

RIPA buffer (see recipe), ice-cold 

Lithium chloride solution: 0.5 M LiCl/0.1 M Tris·Cl, pH 8.0 (see APPENDIX 2A for 

Tris·Cl), ice-cold 

RM×3 (see recipe) 

2 mg/ml myelin basic protein (MBP), or other appropriate phosphorylation 

substrate at appropriate (usually lower) concentration 


7 × 10–cm 15% SDS-PAGE gel with stacking gel (UNIT 6.1; also see Basic Protocol 

1) 

Prestained protein markers 

Staining solution (see recipe) 

Destaining solution (see recipe) 





Probe sonicator (e.g., Branson) 



End-over-end rotator 

30◦C Thermomixer (Eppendorf) or water bath 

Boiling water bath 

Flat container for staining/destaining gel 


and autoradiography or phosphor imaging (UNIT 6.3) 








14.3.13 




14.3.14 


1. Grow Rat1 cells in six 6-cm tissue culture dishes in 4 ml DMEM containing 10% 

FBS to subconfluency (∼0.5 × 10 6 cells/dish). 


2. Serum-starve the Rat1 cells by removing the culture medium from each dish, 

replacing it with 2 ml starvation medium, and incubating 18 hr. 
















appropriate times and the cells harvested within a short period of time (within 5 to 

10 min for all dishes). If the influence of the stimulating agent on the particular cells is 

not yet known, a positive control should be included, such as a dish treated with 50 µl 

peroxovanadate (VOOH) solution (see recipe), for a final concentration of 100 µM sodium 

orthovanadate and 200 µMH2O2, which nonspecifically activates many signaling events 

in most tissue culture cells. 


cells. Rinse the dishes twice with 5 ml ice-cold PBS and once with 5 ml ice-cold 

buffer A. Be sure to remove all of the PBS after the last wash. Place dishes on ice. 



1.5 min. so that all six dishes are harvested within 5 to 10 min. 

5. Add 250 µl of ice-cold buffer H to each dish, tilt the dish gently (preferably on 

ice), and scrape the cells into the buffer using a plastic (or rubber) policeman. Using 

precooled pipet tips, transfer the cells and buffer to prelabeled, precooled 1.5-ml 

plastic test tubes. 

Special consideration should be given to the composition of the buffer H (see Reagents 

and Solutions). The authors recommend using β-glycerophosphate, which serves both 

as a buffer and a general phosphatase inhibitor, rather than Tris or HEPES. Sodium 

orthovanadate is used to inhibit tyrosine phosphatases, and the mixture of pepstatin A, 

aprotinin, leupeptin, and benzamidine is used to inhibit proteinases. This buffer, when 

cold, blocks most of the phosphatase and proteinase activities in cell extracts. 

6. Disrupt the cells by sonication using two 7-sec 50-W pulses with a 20-sec interval 

per ∼0.5-ml sample on ice. 

Over the years, several methods of protein extraction from cells have been successfully 

used in the study of protein kinases. In this protocol, which utilizes sonication, proteins are 

extracted from the cytosolic and nuclear fractions, but not from the membrane fraction, and 

therefore can be considered as cytosolic extract. Cellular extraction with nonionic detergents, 

which extract proteins from membranal, cytosolic, and some nuclear fractions of the 

cell, makes determination of the protein concentration somewhat difficult, but is often used. 

Extraction with RIPA buffer or by freezing-thawing can also be used for some kinases. 


7. Microcentrifuge the cellular extracts 15 min at 14,000 × g, 4 ◦ C. Transfer the resulting 

supernatants (cytosolic extracts), which contain the protein to be examined 

for phosphorylation, to fresh precooled microcentrifuge tubes. Take a 5- to 20-µl 

aliquot of each extract and determine protein concentration (steps 8 to 12). Store the 

remainder of each cytosolic extract on ice until needed for the immunoprecipitation 

and phosphorylation steps (steps 13 to 27). 

The protein concentration of each sample should be determined so that the amounts of 

proteins from the different samples can be compared, making it possible to determine the 

relative amounts of protein kinases in all samples accurately. If samples are compared 

based on cell number, differences of up to 20% in the amount of protein may result. Such 

differences may cause even larger ones in the following steps. 

Determine protein concentration 

8. Dilute 10 µl of each cytosolic extract sample in 190 µl buffer A (a final dilution of 

1:20) in labeled tubes. 






plate. 




of the same microtiter plate. Add 200 µl of the Bradford protein assay reagent to all 

wells. 


at 595 nM. 

12. From the optical densities, calculate the protein concentrations of the samples. 

Prepare antibody-conjugated protein A–Sepharose beads 

13. Place ∼150 µl protein A–Sepharose beads in a 1.5-ml microcentrifuge tube and add 

1 ml of PBS. Let the beads swell for 10 min at room temperature. 

Although protein A–conjugated Sepharose is recommended for this method, other commercially 

available protein A–conjugated resins, such as agarose, HiTrap, etc., may be 

used. Protein G–coupled resins are sometimes required to immunoprecipitate certain 

types of monoclonal antibodies (Table 7.2.1). If resins are supplied as ready-to-use suspensions, 

this swelling step is not necessary. 

14. Microcentrifuge the swollen beads 1 min at 14,000 × g, room temperature, and 

remove the supernatant. Wash the swollen beads three times, each time by adding 

1 ml PBS, centrifuging again as before, and discarding the supernatant. 

15. Combine 180 µl of the swollen packed beads with 320 µl PBS. Add the recommended 

amount of the antibody for immunoprecipitation (e.g., ∼10 µg anti-ERK C terminus). 

Rotate the mixture 1 hr at room temperature on an end-over-end rotator to allow the 

antibodies to bind to the protein A. 

Alternatively, this step can be done at 4 ◦ C for 16 hr. 

Usually, anti-C-terminal antibodies are used for the determination of kinase activity 

because their binding to the kinase does not interfere with the kinase activity. Ideally, 

the volumes listed here should be sufficient for ten reactions, but, because of the density 

of the beads, they will probably only be sufficient for eight reactions. Depending on the 



14.3.15 




14.3.16 

number of reactions to be performed, the amounts given here can be scaled up as long as 

the proportions are maintained. 

For easy handling of the resin, the ends of the pipet tips can be cut to enlarge their 

openings. 

16. Microcentrifuge the antibody-conjugated beads 1 min at 14,000 × g, room temperature. 

Resuspend the beads in 1 ml PBS, then centrifuge again as before and 

remove the supernatant. Wash three times, each time by adding 1 ml ice-cold PBS, 

centrifuging again as before, and removing the supenatant. Resuspend the washed 

beads in an equal volume of ice-cold buffer A (∼250 µl for∼250 µl of beads). 

Either use the antibody-conjugated beads immediately, or store at 4 ◦ C until used. It is 

best to use the conjugated beads within 3 days of preparation. 

17. In precooled 1.5-ml microcentrifuge tubes, add 30 µl of the antibody-conjugated 

bead suspension from step 16 (equivalent to 15 µl packed beads) to 300 µl of 

cytosolic extract (from step 7) containing 50 to 500 µg total protein (determined as 

in steps 8 to 12) in buffer H. Rotate on an end-over-end rotator for 2 hr at 4 ◦ C. 

Several immunoprecipitation methods have been developed. These methods usually vary 

with respect to the order in which the antibodies and protein A are added to the cell 

extracts. In the protocol described here, the antibodies are conjugated to protein A beads 

and only then added to the cytosolic extracts. This procedure minimizes the time the 

samples are incubated with the antibodies, and thereby minimizes exposure of the desired 

kinases to phosphatases and proteinases in the extracts. Furthermore, this procedure 

ensures that only antibodies recognized by protein A will be used for the immunoprecipitation. 

Antibodies that are not recognized by protein A can bind to the desired antigen, but 

they will not be precipitated when protein A beads are added; therefore, such antibodies 

will reduce the efficiency of immunoprecipitation. 

18. Centrifuge the incubation mixture 1 min at 14,000 × g,4 ◦ C. Remove and discard the 

incubation supernatant from the antibody-conjugated beads. Add 1 ml ice-cold RIPA 

buffer, centrifuge again as before, and discard the supernatant. Wash the pellet twice, 

each time by adding 1 ml ice-cold 0.5 M lithium chloride solution, centrifuging again 

as before, and removing the supernatant, then wash twice, each time by adding 1 ml 

ice-cold buffer A, centrifuging again, and discarding the supernatant. 

These stringent washes are important because they will remove most sticky protein kinases 

that might nonspecifically interact with the protein A beads. 

Perform phosphorylation reaction 

19. After the last wash step in 18, completely remove buffer A from the conjugated beads 

by microcentrifuging again after the buffer has been aspirated from above the pellet 

(without resuspension), then gently removing the residual buffer above the beads. 

20. Resuspend the bead pellet in 15 µl water. 

At this stage, prepare the bench for working with small amounts of radioactivity (see 

APPENDIX 1D). 

21. Add 10 µlofRM×3 to each tube. 

See Reagents and Solutions for a discussion of the components of RM×3. 

22. Start the phosphorylation reaction by adding 5 µl of the phosphorylation substrate 

(e.g., 2 mg/ml MBP) to the tube and placing the mixture in an Eppendorf Thermomixer 

(or water bath) at 30 ◦ C. Incubate 20 min at 30 ◦ C with either constant or 

frequent shaking. 

Although MBP is probably not a physiological substrate for any MAPK, it is a good 

general substrate for many kinases, including ERKs, in vitro. Substrates should be well 

phosphorylated by the desired kinases to allow accurate detection of the phosphorylation 


kinetics. Therefore known substrates of the MAPK can be used as good substrates instead 

of MBP, but those are usually used at lower concentrations in the phosphorylation 

reaction. 

23. End the phosphorylation reaction by adding 10 µlof4× sample buffer to each tube. 

24. Place samples in a boiling water bath for 5 min. Microcentrifuge 1 min at 14,000 × g, 

room temperature. Load the resulting supernatants on a 7 × 10–cm, 15% SDS-PAGE 

gel with stacking gel (UNIT 6.1). Load prestained markers on gel as well. Run the gel 

at 150 V (constant voltage) for ∼1 hr or until the dye front is 0.5 cm from the bottom 

(UNIT 6.1). 

Usually, prestained markers are loaded into the first or second lane of the gel so that the 

first lane can be located on the dried gel and the molecular weight of the detected proteins 

determined. 

Since determination of enzymatic activity is not always accurate when enzymes (in this 

case protein kinases) are bound to beads, the kinase(s) of interest can be released from 

the beads at step 19 or 20 by using excess immunizing peptide. The phosphorylation 

reaction (steps 21 to 23) can thus be performed without the interference of the beads. 

Then the activity can be detected as described (step 27) or by a paper assay. For the 

paper assay, the phosphorylation reaction is terminated by spotting 20 µl of each sample 

on phosphocellulose paper squares (Whatman P81), which are immediately washed with 

150 mM phosphoric acid. Phosphate incorporation is then measured using a scintillation 

cocktail. Gel quantitation can also be carried out by densitometry after imaging the gel 

as in step 27. 

25. Turn the power supply off and disconnect the leads. Remove the glass plates containing 

the gel and remove the gel from the plate. 

26. Place the gel in a flat container and stain the gel with 50 ml staining solution for 

20 min at room temperature. Destain with four 30-min changes of 50 ml destaining 

solution. 

This extensive destaining removes excess free [γ - 32 P]ATP, which would affect the background. 

In some cases, the extensive destaining is not necessary, or the proteins can be 

transferred to a nitrocellulose membrane and then be imaged as described below. 

27. Place the gel on a sheet of Whatman 3MM paper, cover the gel with a clear plastic 

wrap, and dry the gel in a gel dryer (usually 1.5 hr at 80 ◦ C is sufficient). Expose the 

gel in a phosphor imager or autoradiograph on X-ray film (UNIT 6.3). 

Bands should appear at 16 to 21 kDa, corresponding to the molecular weight of the four 

MBP isoforms. 

IN-GEL KINASE ASSAY 

If the identity of the kinase is not known or there are no specific antibodies available 

for the kinase, the in-gel kinase assay may be used instead of the immunoprecipitation 

protocol (Basic Protocol 2). This in-gel protocol involves copolymerization of a substrate 

with the polyacrylamide gel followed by electrophoresis of the protein sample(s) on the 

gel. After several rounds of denaturation and renaturation, a phosphorylation reaction is 

performed with the gel, and the phosphorylated bands are visualized by autoradiography 

or phosphor imaging. With this method, the molecular weight of the protein kinase is 

revealed and unknown protein kinases can be identified (the molecular weight of most 

MAPK components is shown in Table 14.3.1). However, not all protein kinases can be 

renatured under the conditions of this protocol, and the linear range of this assay is usually 

limited. Therefore, this method should not be routinely used to monitor and characterize 

known protein kinases. 

BASIC 

PROTOCOL 3 



14.3.17 




14.3.18 

CAUTION: When working with radioactive materials, take appropriate precautions to 

avoid contamination of the experimenter and the surroundings. Carry out the experiment 

and dispose of wastes in an appropriately designated area, following guidelines provided 

by the institutional Radiation Safety Officer (also see UNIT 7.1 and APPENDIX 1D). 

Materials 





50 µg/ml epidermal growth factor (EGF) in EGF buffer 

EGF buffer: 0.5 mg/ml bovine serum albumin (BSA) in phosphate buffered saline 




Buffer H (see recipe) containing 1% (v/v) Triton X-100, ice-cold 


Protein standards: 5, 10, 20, 50, 100 and 200 µg/ml BSA in Buffer H (see recipe 

for buffer H) 





30% acrylamide/0.8% bisacrylamide (Table 14.3.1) 

10% (w/v) ammonium persulfate (prepare fresh) 

Tetramethylethelendiamine (TEMED) 

2 mg/ml myelin basic protein (MBP) 


Running buffer (see recipe) 

20% (v/v) isopropanol/50 mM HEPES, pH 7.6 

Renaturation buffer: 50 mM HEPES, pH 7.6 containing 5 mM 2-mercaptoethanol 

Renaturation buffer containing 6 M urea 

Renaturation buffer containing 0.05% (v/v) Tween 20 

In-gel kinase buffer (see recipe) 

In-gel kinase reaction solution (see recipe) 

5% (w/v) trichloroacetic acid (TCA)/1% (w/v) sodium pyrophosphate 







Gel-casting apparatus: 7 × 10–cm glass plates, 1.5-mm spacers, and 1.5-mm comb 

with 10 teeth (also see UNIT 6.1) 

Gel electrophoresis apparatus and power supply (also see UNIT 6.1) 

Flat containers for washing gel 

30 ◦ C water bath with proper shielding for radioactive work 


and autoradiography (UNIT 6.3) 








1. Grow Rat1 cells in six 6-cm tissue culture dishes in DMEM containing 10% FBS to 

subconfluency (∼0.5 × 10 6 cells/dish). 


2. Serum-starve the Rat1 cells by removing the culture medium from each dish, replacing 

it with 2 ml starvation medium, and incubating 18 hr. 
















appropriate times and the cells harvested within a short period of time (within 5 to 10 

min for all dishes). If the influence of the stimulating agent on the particular cells is 

not yet known, a positive control should be included, such as a dish treated with 50 

µl peroxovanadate (VOOH) solution (see recipe), for a final concentration of 100 µM 

sodium orthovanadate and 200 µMH2O2, which nonspecifically activates many signaling 

events in most tissue culture cells. 


cells. Rinse the dishes twice with 5 ml ice-cold PBS and once with 5 ml ice-cold 

buffer A. Be sure to remove all of the PBS after the last wash. Place dishes on ice. 



1.5 min, so that all six dishes are harvested within 5 to 10 min. 

5. Harvest cells in 250 µl buffer H containing 1% Triton X-100 on ice, scraping the 

dishes with a plastic or rubber policeman and transferring the homogenate to a 1.5ml 

microcentrifuge tube. Microcentrifuge 15 min at 14,000 × g, 4 ◦ C. Collect the 

supernatant and transfer to a new 1.5-ml tube. 

For this protocol, extraction with detergent is usually better than sonication. 

Determine protein concentration 

6. Dilute 10 µl of each extract from step 5 in 190 µl buffer A (a final dilution of 1:20) 

in labeled tubes. 






14.3.19 




14.3.20 



plate. 




of the same microtiter plate. Add 200 µl of Bradford protein assay reagent to all wells. 


at 595 nM. 

10. From the optical densities, calculate the protein concentrations of the samples. Take 

an aliquot of each extract corresponding to 50 to 100 µg of protein and place in a 

new 1.5-ml microcentrifuge tube. 

11. Add 1/3 vol of 4× sample buffer to each tube. Keep at 4 ◦ C, without boiling. 

Electrophorese the sample 

12. Assemble 7 × 10–cm glass plates and 1.5-mm spacers for casting the polyacrylamide 

gel. 

13. Prepare a 12% separating gel containing MBP by mixing the following (total volume, 

8 ml): 

0.7 ml H2O 

2.0 ml 2 mg/ml MBP 

2.0ml1.5MTris·Cl, pH 8.8 

3.2 ml 30% acrylamide/0.8% bisacrylamide 

100 µl 10% ammonium persulfate 

6 µl TEMED. 

14. Pour the separating gel (UNIT 6.1). Overlay the top of the separating gel with water. 

Allow the gel to polymerize ∼30 to 45 min. 

15. Prepare 3% polyacrylamide stacking gel by mixing the following (5 ml total): 

550 µl 30% acrylamide/0.8% bisacrylaimde solution 

625 µl 0.5MTris·Cl, pH 6.8 

3.7 ml H2O 

120 µl ammonium persulfate 

5 µl TEMED 

5mltotal. 

16. Remove the water from the top of the polymerized separating gel. Using a Pasteur 

pipet, layer ∼2 ml of the stacking gel on top of the separating gel (UNIT 6.1) to fill up the 

apparatus. Insert comb into the stacking gel and allow it to polymerize 10 to 20 min. 

17. Place the polymerized gel in the electrophoresis apparatus, add running buffer, and 

check for leaks. 

18. Load ∼40 µl of the samples (from step 13; containing ∼70 µg protein per lane) and 

10 µl of the prestained protein markers (according to the manufacturer’s instructions) 

onto the gel. Electrophorese at 100 V for ∼1 hr or until the dye front is ∼0.5 cm 

from the bottom (UNIT 6.1). 

The gel should not be heated above 30 ◦ C, therefore, the voltage used for electrophoresis 

should not be higher than 100 V. 

19. When electrophoresis is completed, turn the power supply off, remove the glass 

plates from the electrophoresis apparatus, and remove the gel from the plates. 


Wash the gel and renature the proteins 

20. Cut off the stacking gel. Place the separating gel in a flat container and wash twice, 

each time with 100 ml of 20% isopropanol/50 mM HEPES, pH 7.6, for 30 min at 

room temperature with shaking on a platform shaker. Wash the gel twice more, each 

time with 100 ml renaturation buffer for 30 min at room temperature with shaking, 

then wash the gel twice more, each time with 100 ml of 6 M urea (in renaturation 

buffer) for 15 min with shaking at room temperature. Leave the last urea wash on 

the gel and proceed to step 21. 

If necessary, the second wash with 20% isopropanol/50 mM HEPES can be done overnight 

at 4 ◦ C. 

21. Place the gel in a 4 ◦ C cold room, remove 50 ml of the 6 M urea, add 50 ml renaturation 

buffer containing 0.05% Tween 20, and shake for 15 min on a platform shaker. 

The washing solution is now 3 M urea in renaturation buffer containing Tween 20. 

22. Remove 50 ml of the washing solution, add 50 ml of renaturation buffer containing 

0.05% Tween 20, and shake for additional 15 min 

This reduces the urea to 1.5 M. 

23. Remove 50 ml of the washing solution once more and replace with renaturation 

buffer/0.05% Tween 20, so that the washing solution contains 0.75 M urea. Shake 

for 15 min. 

24. Wash the gel twice, each time with 100 ml renaturation buffer containing 0.05% 

Tween 20 for 15 min with shaking at 4 ◦ C. Wash a third time with the same solution 

with shaking overnight in the cold room. 

Detect kinase activity 

25. Remove the washing buffer and incubate the gel in 30 ml in-gel kinase buffer for 30 

min at 30 ◦ C. Remove the buffer and add 20 ml of in-gel kinase reaction solution. 

At this stage, the amount of radioactive material is very high, and therefore the reaction 

should be performed with a proper shielding. Make sure that the gel is straight in the 

flat container. Unequal distribution of the phosphorylation buffer may lead to wrong 

concentrations of the ingredients and thereby interfere with the phosphorylation reaction. 

26. Wash gel carefully four times, each time with 3 ml 5% TCA/1% sodium pyrophosphate 

for 15 min at room temperature. 

If the gel is still very radioactive, continue washing overnight. 

27. Dry the gel and subject to autoradiography (UNIT 6.3). 

Bands should appear where kinases are present and cause phosphorylation of the MBP 

copolymerized in the gel (see Fig. 14.3.2). 

REAGENTS AND SOLUTIONS 

Use deionized or distilled water in all recipes and protocol steps. For common stock solutions, 

see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX. 

Buffer A 

50 mM β-glycerophosphate, pH 7.3 

1.5 mM EGTA 

1.0 mM EDTA 

1.0 mM dithiothreitol (DTT) 

0.1 mM sodium orthovanadate 

Store up to 2 weeks at 4◦C Recipe from Ahn et al. (1990). 



14.3.21 




14.3.22 

Buffer H 


1.5 mM EGTA 

1.0 mM EDTA 


Store solution with above components up to 3 months at 4◦C Immediately before use, add: 

1.0 mM DTT 

1.0 mM benzamidine 

10 µg/ml aprotinin 

10 µg/ml leupeptin 

2.0 µg/ml pepstatin A 

Recipe from Seger et al. (1992). “H” refers to homogenization buffer. 

Destaining solution 

15% (v/v) isopropanol 

7% (v/v) acetic acid 

Store up to 1 month at room temperature 

In-gel kinase buffer 

20 mM HEPES, pH 7.6 

20 mM MgCl2 

Store up to 1 month at 4◦C In-gel kinase reaction solution 


20 mM MgCl2 

2mMDTT 

20 µM ATP (unlabeled) 

100 µCi [γ-32P]ATP Store up to 2 weeks at −20◦C Peroxovanadate (VOOH) solution 

Mix 200 µl of 20 mM sodium orthovanadate with 800 µl of10mMH2O2 in PBS 

(see APPENDIX 2A for PBS). Allow mixture to incubate 15 min at room temperature 

before use. Prepare fresh. 

RIPA buffer 

137 mM NaCl 

20 mM Tris·Cl, pH 7.4 (APPENDIX 2A) 

10% (v/v) glycerol 

1% (v/v) Triton X-100 

0.5% (w/v) sodium deoxycholate 

0.1% (w/v) SDS 

2.0 mM EDTA 

Store with above components up to 1 month at −20◦C 1.0 mM phenylmethylsulfonyl fluoride (PMSF, add fresh) 

20 µM leupeptin 

RM×3 


30 mM MgCl2 

100 µM [γ-32P]ATP (∼4000 cpm/pmol) 

0.9% (w/v) bovine serum albumin (BSA) 

continued 


3mMDTT 

3mMEGTA 


Store up to 3 month at −20 ◦ C 

The most important components of the reaction mixture (also see Basic Protocol 2, step 

21) are the Mg 2+ and [γ - 32 P]ATP, which are essential for the phosphorylation reaction. 

The authors recommend the use of 100 µM ATP with ∼4000 cpm/pmol of the labeled ATP, 

which provides an extended linear range and reproducible results. When the enzymatic 

activity of the kinases is very low, which makes detection of phosphorylation difficult, the 

concentration of cold ATP should be reduced to 10 to 20 µM and the amount of radioactive 

material elevated. Addition of labeled ATP alone is not recommended because this will 

result in a nanomolar concentration of ATP, which is much below the Km for ATP and 

may lead to nonspecific phosphorylation. The β-glycerophosphate in the reaction mixture 

serves as a buffer, but it can also inhibit residual phosphatases that may have nonspecifically 

bound to the beads. The BSA serves as a carrier protein, but when purity is required, it can 

be eliminated. The EGTA chelates Ca 2+ , which may interfere with some kinase activities, 

DTT keeps the proteins reduced, and sodium orthovanadate inhibits tyrosine phosphatases. 

Running buffer 


188 mM glycine 

0.1% (w/v) SDS 


Sample buffer, 4× 



8% (w/v) SDS 

8% (v/v) β-mercaptoethanol 

0.2% (w/v) bromphenol blue 

Store up to 3 months at −20 ◦ C 

Staining solution 

40% (v/v) methanol 

7% (v/v) acetic acid 


Store up to 3 months at room temperature 

Transfer buffer 


50 mM glycine 


Tris-buffered saline with Tween 20 (TBST) 


150 mM NaCl 

0.05% (v/v) Tween 20 


COMMENTARY 

Background Information 

Sequential activation of kinases (protein 

kinase cascades) is a common mechanism of 

signal transduction in many cellular processes 

(Seger and Krebs, 1995). The best studied protein 

kinase cascades known today are mitogenactivated 

protein kinase (MAPK) signaling 

cascades, which are the topic of this unit (Seger 



14.3.23 




14.3.24 

and Krebs, 1995; Chen et al., 2001; Morrison 

and Davis, 2003; Raman and Cobb, 2003; 

Rubinfeld and Seger, 2004). Each of these signaling 

cascades consists of up to five tiers 

of protein kinases that sequentially activate 

each other by phosphorylation. The similarity 

among the enzymes that comprise each tier 

in the various cascades categorizes them into 

a superfamily of kinases with similar structure 

and function. 

The activation of each of these cascades 

is initiated either by small GTP-binding proteins 

or by adaptor proteins that transmit 

the signal to protein kinases, commonly referred 

to as MAPK kinase kinases (MAP3Ks). 

In some cases, the activation is mediated 

by upstream, Ste20-like kinases, which are 

termed MAP4Ks, although in many cases the 

MAP3Ks can be activated directly by the GTP 

binding proteins, or the MAP4Ks bypass the 

MAP3Ks (Fig. 14.3.1, Table 14.3.1). The signal 

is then transmitted downstream in the cascade 

by the sequential activation of MAPK kinase 

(MAPKK), MAPK and MAPK-activated 

protein kinases (MAPKAPKs), and, in a few 

cases, also kinases downstream of this tier. The 

existence of as many as three to six tiers in each 

of the MAPK cascades is probably essential 

for signal amplification, specificity determination, 

and tight regulation of the transmitted 

signal. 

The four distinct MAPK cascades that are 

currently known are named according to the 

subgroup of their MAPK components. These 

are (1) the extracellular signal-regulated kinase 

(ERK) cascade; (2) the c-Jun N-terminal 

kinase [JNK; or stress activated protein kinase 

1 (SAPK1) cascade]; (3) the p38 cascade 

(Freshney et al., 1994; Han et al., 1994; 

Lee et al., 1994); and (4) the BMK1 (ERK5) 

cascade (see Seger and Krebs, 1995 and 

Chuderland and Seger, 2005, for reviews). The 

identification of ERK7/8 (Abe et al., 1999) 

indicates that additional full MAPK cascades 

can emerge. The ERK cascade seems to participate 

mainly in the transmission of mitogenic 

signals, whereas the p38 and JNK cascades 

transmit mainly stress signals. BMK seems to 

be activated equally by mitogens and stress 

stimuli. These MAPK cascades cooperate to 

transmit signals to their intracellular targets, 

and thus to initiate cellular processes such 

as proliferation, differentiation, development, 

stress response, and apoptosis. In this section, 

the various MAPK cascades will be briefly reviewed. 

The ERK cascade, also known as the p42, 

p44 MAPK cascade, was the first MAPK 

cascade elucidated (Seger and Krebs, 1995). 

This cascade is initiated in many cases by 

the small G-protein Ras, which, upon stimulation, 

causes membranal translocation and activation 

of the protein serine/threonine kinase, 

Raf1 (reviewed in Hagemann and Rapp, 1999). 

Once activated, Raf1 continues the transmission 

of the signal by phosphorylating two regulatory 

serine residues located in the activation 

loop of MEK, thus causing its full activation 

(Kyriakis et al., 1992). Interestingly, Raf-1 activation 

can be facilitated by PKC, and this can 

implicate the PKC as a MAP4K of the ERK 

cascade. Other kinases that act in the MAP3K 

level of the cascade under various conditions 

are—(1) B-Raf (Peraldi et al., 1995), which is 

usually activated by the small G-protein Rap1 

(Hagemann and Rapp, 1999) in combination 

with phosphorylation by MLK3 (Chadee and 

Kyriakis, 2004); (2) A-Raf (Hagemann and 

Rapp, 1999); (3) Mos, which acts specifically 

in the reproductive system (Gotoh and 

Nishida, 1995); (4) the protooncogene TPL2 

(Salmeron et al., 1996); (5) under stress conditions, 

possibly also MEKK1 (Lange-Carter 

et al., 1993) and other MAP3Ks such as MLK2 

(Hirai et al., 1997), MLTK, (Gotoh et al., 

2001), MLK3 (Chadee and Kyriakis, 2004), 

and even IRAK (MacGillivray et al., 2000), although 

the direct effect of these kinases is still 

controversial. Another kinase on this level is 

kinase suppressor of Ras (KSR), which seems 

to act mainly as a scaffold protein for some of 

the MAP3K and MAPKK components of the 

cascade (Morrison and Davis, 2003); but the 

role of its catalytic activity is still controversial 

(Kolesnick and Xing, 2004). In any case, all 

the indicated MAP3Ks seem to phosphorylate 

the same regulatory residues of MEK, which 

are required for its full activation (Ahn et al., 

1991; Seger et al., 1992). Activated MEK is a 

dual-specificity protein kinase, which appears 

to be the only kinase capable of specifically 

phosphorylating and activating the next kinase 

in this cascade, i.e., ERK (Boulton et al., 1990, 

1991). ERK activation requires phosphorylation 

of two regulatory residues, threonine and 

tyrosine, which reside in a TEY phosphorylation 

motif (Payne et al., 1991; Canagarajah 

et al., 1997). Although phosphorylation of 

threonine and tyrosine residues is essential for 

the activation of all MAPKs, in the other cascades, 

the identity of the middle amino acid 

in the TXY motif of the MAPK is different, 


and this determines the specificity of the signal 

(Seger and Krebs, 1995). 

ERK appears to be an important regulatory 

molecule, which, by itself, can phosphorylate 

regulatory targets in the cytosol such 

as phospholipase A2 (PLA2; Lin et al., 1993). 

It can translocate into and phosphorylate nuclear 

substrates such as the transcription factors 

Elk1 (Gille et al., 1992), cFos (Murphy 

et al., 2002), p53 (Milne et al., 1994), ERF 

(Sgouras et al., 1995), PEA-3 (O’Hagan et al., 

1996), ERM (Janknecht et al., 1996), ER81 

(Janknecht et al., 1996), Ets1/2 (Yang et al., 

1996), SAP-1a (Strahl et al., 1996), and cJun 

(Morton et al., 2003), or it can transmit the signal 

to the MAPKAPK level. The main MAP- 

KAPK of the ERK cascade is RSK (Sturgill 

et al., 1988), which can also translocate to the 

nucleus upon activation and phosphorylate a 

set of nuclear substrates different from those 

phosphorylated by ERK. Additional MAP- 

KAPKs are the mitogen- and stress-activated 

kinase (MSK; Deak et al., 1998) and the 

MAPK-interacting kinases (MNKs; Fukunaga 

and Hunter, 1997; Waskiewicz et al., 1997), 

which are activated equally by the related p38 

cascade. Finally, the MAPKAPK3 (McLaughlin 

et al., 1996) and MK5 (Ni et al., 1998) 

cascades are only slightly related to the ERK 

cascade and are mainly activated by the p38 

cascade (for review see Roux and Blenis, 

2004). MAPKAPKs can regulate the activity 

of additional kinases (e.g., Myt1; Palmer et al., 

1998), but those are usually not considered to 

be genuine members of the MAPK cascade. 

ERKs are activated primarily by mitogenic 

signals, whereas other MAPK cascades (p38, 

JNK) are activated mainly by cellular stresses 

such as heat shock, ischemia, UV irradiation, 

and cytokines (Davis, 2000); therefore they 

are referred to as stress-activated protein kinase 

(SAPK) cascades (Kyriakis and Avruch, 

1996). The p38 cascade consists of MAPKs 

that contain a glycine residue in their activation, 

TXY motif (TGY; Han et al., 1994). 

Many kinases on the MAP3K and MAP4K 

levels have been implicated in the p38 cascade 

(Fig. 14.3.1, Table 14.3.1); however, their individual 

roles are not yet fully elucidated. Moreover, 

in spite of their sequence similarity to 

the Ste20 and Ste11 kinases (the MAP4K and 

MAP3K in Saccharomyces cerevisiae that operate 

in a sequential order; Herskowitz, 1995), 

the mammalian kinases do not always act in a 

sequential order, and may use different mechanisms 

to activate the rest of the cascade (Dan 

et al., 2001; Hagemann and Blank, 2001). 

Independent of the particular MAP3K or 

MAP4K, the signals of the p38 cascade are 

funneled into two main MAPKKs: MKK3 

(also known as MEK3 or p38MAPKK; Derijard 

et al., 1995) and MKK6 (also known as 

MEK6, or SAPKK3; Han et al., 1996). Similarly 

to MEK1/2, these are dual-specificity 

protein kinases that phosphorylate the regulatory 

threonine and tyrosine residues of the 

p38s to fully activate them. Although these 

two MKKs are unique in their ability to activate 

p38s and no other MAPKs, the p38s 

seem to be activated by MKK4 (Derijard et al., 

1995) and to some extent also by MKK7 

(Dashti et al., 2001), which are the main activators 

of the JNK cascade (as indicated below). 

The MAPK-level components of this 

cascade are the four p38s (p38α to δ, also 

known as SAPK2a SAPK2b, SAPK3, and 

SAPK4; Han et al., 1994; Goedert et al., 1997), 

which are normally expressed as four main 

proteins of 41, 43, 47, and 51 kDa, and at 

least six additional alternatively spliced forms 

with molecular masses that range between 32 

and 65 kDa. Once these p38s are activated, 

they phosphorylate and modulate the activity 

of a large number of substrates in various 

compartments in the cells (Davis, 2000). 

Thus, the p38s can phosphorylate and activate 

various MAPKAPK-level components including 

MAPKAPK2 and 3 (Stokoe et al., 1992; 

McLaughlin et al., 1996), MSK (Deak et al., 

1998), MNK (Fukunaga and Hunter, 1997; 

Waskiewicz et al., 1997), and MK5 (Ni et al., 

1998). Except for MAPKAPK2, all MAP- 

KAPKs can also be activated by the ERK cascade 

(see above) and therefore present a point 

of cross-talk between the two cascades (for 

review see Roux and Blenis, 2004). Another 

important group of p38 substrates are various 

transcription factors that are usually activated 

under stress conditions, including ATF2, Elk1 

(Raingeaud et al., 1996), CHOP (Wang and 

Ron, 1996), MEF2C (Han et al., 1997), SAP-1 

(Cuenda et al., 1997), and p53 (Huang et al., 

1999). 

Other stress-activated MAPKs are the c-Jun 

amino-terminal kinases (JNKs; also named 

SAPK1), which comprise a third MAPK subgroup 

(Derijard et al., 1994; Kyriakis et al., 

1994). These enzymes are distinct from the 

p38s mainly because they contain a TPY rather 

than a TGY motif in their activation loop. As 

the other MAPK cascades, the JNK cascade 

is triggered by small GTPases (Crespo et al., 

1997), i.e., Rac and CDC42, or it can be activated 

by various adaptor molecules (Davis, 



14.3.25 




14.3.26 

2000). Next, the signals are transmitted via 

MAP4K and MAP3K components that are 

mostly shared with the p38 cascades (Table 

14.3.1). Since the p38 and JNK cascades are 

not always simultaneously activated, the signals 

must be separately regulated to allow separate 

cascades, and this is mostly executed by a 

physical segragation between the components 

by specialized scaffold proteins (Morrison and 

Davis, 2003). At the MAPKK level, the JNKs 

can be activated by MKK4 (MEK4, SEK1, 

JNKK1; Sanchez et al., 1994; Yan et al., 1994), 

and MKK7 (MEK7, JNKK2; Tournier et al., 

1997). These MKKs are able to phosphorylate 

and activate the components in the MAPK 

level, which are JNK1-3 (SAPKs) of molecular 

weights 46, 54, and 52 kDa, respectively 

(Derijard et al., 1994; Kyriakis et al., 1994). 

No enzymes in the MAPKAPK level have 

been identified for JNKs, which appear to be 

major regulators of nuclear processes, in particular, 

transcription. Shortly after activation, 

JNKs translocate to the nucleus where they 

physically associate with and activate their targets, 

mostly transcription factors such as c-Jun 

(Hibi et al., 1993), ATF2 (Gupta et al., 1995), 

Elk1 (Whitmarsh et al., 1995), p53 (Milne 

et al., 1995), JUN-D (Kallunki et al., 1996), 

PEA3 (O’Hagan et al., 1996), NFAT4 (Chow 

et al., 1997), c-Myc (Noguchi et al., 1999), 

and SAP-2/Net (Ducret et al., 2000). Interestingly, 

the JNKs also participate in the induction 

of apoptosis by several proapoptotic 

drugs, and this is mediated by the phosphorylation 

of various death-related components in 

various cellular compartments (for review see 

Varfolomeev and Ashkenazi, 2004). 

Another MAPK subgroup is the BMK (also 

known as ERK5; Zhou et al., 1995; Abe et al., 

1996), with a molecular weight of 110 kDa. 

Like ERK1/2, BMK contains a TEY phosphorylation 

motif in its activation loop, but because 

it is not activated at all by MEK1 and 

2, it forms a separate signaling cascade. BMK 

seems to be activated equally by stress (Zhou 

et al., 1995; Abe et al., 1996) and by mitogenic 

signals (Kato et al., 1998), and thereby 

participates in the regulation of a variety of 

cellular processes including oncogenic transformation 

(English et al., 1999). The cascade 

is thought to be activated by the adaptor protein 

Lad, which operates downstream of c-Src, 

at least in the pathway that leads to BMK activation 

by growth factors (Sun et al., 2003). In 

turn, Lad transmits the signal to protein kinases 

on the MAP3K level of the cascade, MEKK2 

and MEKK3, which are implicated also in the 

activation of the p38 and JNK cascade (Chao 

et al., 1999; Chayama et al., 2001). The signal 

is then transmitted downstream the cascade to 

the MAPKK, MEK5 (Zhou et al., 1995), which 

specifically phosphorylates and activates the 

kinase on the MAPK level, BMK. Interestingly, 

MEK5 and BMK are often localized in 

the cell nucleus (Raviv et al., 2004), where 

BMK associates with several transcription factors 

substrates including c-Myc (English et al., 

1998), MEF2 family members (Kato et al., 

1997; Yang et al., 1998), c-Fos (Kamakura 

et al., 1999), SAP1a (Kamakura et al., 1999), 

Fra-1 (Terasawa et al., 2003), and possibly 

also NFκB (Pearson et al., 2001). Importantly, 

BMK also influences transcription through a 

direct transcriptional activity, as shown in the 

activation of the Nur77 gene upon calcium signals 

in T cells (Kasler et al., 2000). In addition 

to its nuclear activity, BMK mediates several 

cytosolic functions including phosphorylation 

of connexin-43 (Cameron et al., 2003) and 

of the serum- and glucocorticoid-inducible kinase 

SGK (Hayashi et al., 2001), implicating 

the latter as a MAPKAPK in the ERK5 cascade. 

Besides these four MAPK cascades, which 

are composed of similar components in each 

level, other protein kinases and kinase cascades 

are activated in response to mitogenic 

stimulation. First, two MAPKs termed ERK7 

and ERK8, which are similar to each other 

(Abe et al., 2001), have been identified. Although 

their mode of activation is not clear, 

they are likely to operate within an independent, 

fifth MAPK cascade. Another cascade is 

that of ERK3, which, in spite of its 50% identity 

to ERK1/2, is not considered a MAPK 

because of the absence of the TEY motif 

in its activation loop (Boulton et al., 1991). 

ERK3 is activated by an ERK3K with an unknown 

identity (Cheng et al., 1996), and it 

seems to function mainly in muscles. In addition, 

the NIK-IKK (Malinin et al., 1997), 

PI3K-PDK-AKT-GSK3 (Cohen et al., 1997), 

Rho-dependent pathways (Leung et al., 1995), 

and the PKA-phosphorylase kinase pathway 

(Brushia and Walsh, 1999) operate in a kinase 

cascade. However, because of their distinct 

characteristics, these pathways are not 

considered MAPK cascades, although they are 

involved in transmission of mitogenic signals. 

The inactivation of many components of the 

MAPK cascade is mediated by a large number 

of distinct phosphatases, including MKP, 

PTPs, and PPs (for review see Yao and Seger, 

2004). 


All the pathways mentioned are apparently 

activated to some extent by distinct extracellular 

agents, and as a result of their action 

in an elaborate network, determine the outcome 

of each stimulation. The full dimensions 

of this network, the mode of regulation 

of its components, and the mechanism by 

which these cascades determine cell fates in response 

to various stimuli, have yet to be fully 

elucidated. 

Critical Parameters 

Several points should be considered when 

using Basic Protocol 2 (immunoprecipitation 

followed by phosphorylation). One of the most 

important parameters for the success of this 

protocol depends on the method of protein 

extraction used. Since the MAPKs are localized 

within cells, the cellular membranes 

must be disrupted to access the desired targets. 

The protein kinases of interest must then 

be obtained and preserved in their active form, 

while decreasing the amount of nonrelevant 

kinases. For example, activated Raf-1 can be 

present in Golgi membranes, which might not 

be disrupted by some extraction procedures, 

but are disrupted if RIPA buffer is used for 

extraction. The method for cellular extraction 

described in Basic Protocol 1 can be effectively 

used for detection of most MAPKs by 

immunoprecipitation or other methods. Sonication 

disrupts the plasma membrane but does 

not solubilize it, and therefore the resulting extracts 

(referred to as cytosolic extracts) should 

contain both cytosolic and some nuclear fractions. 

Depending on the subcellular localization 

of the proteins of interest, other extraction 

methods can be applied. For example, cellular 

extracts obtained with Triton X-100 usually 

contain membrane, cytosolic, and some 

nuclear components, whereas cellular extracts 

obtained with RIPA buffer should contain solubilized 

proteins from most cellular compartment. 

Cellular extraction by addition of hot 

SDS-PAGE sample buffer to cells is not recommended 

because it frees chromatin, which 

causes formation of a gel that is hard to handle. 

Extraction by freeze-thawing is not recommended 

either, because of molecular degradations 

that might occur during the thawing 

phases. 

Another consideration for successful detection 

of phosphoproteins is minimization 

of protein degradation and dephosphorylation. 

During extraction, most cellular organelles 

break, and thus expose phosphoproteins to 

phosphatases and proteinases. Addition of 

specific inhibitors of phosphatases and proteinases 

to the extraction buffers, and extraction 

at low temperatures, minimizes the effect 

of these enzymes. However, since phosphatases 

are usually efficient enzymes, even if 

these precautions are taken extractions should 

be performed as quickly as possible. 

One of the most critical elements for the 

success of the basic methods for immunological 

monitoring of phosphorylation is the quality 

and specificity of the antibodies used. The 

antibodies employed should (1) recognize the 

phosphorylated protein but not its nonphosphorylated 

counterpart, and (2) recognize only 

the desired phosphorylated amino acid, not additional 

phosphorylated sites in either the same 

or different proteins. In addition, the amount 

of proteins in the different samples subjected 

to immunoblot analysis and the dilution of 

the antibodies should be optimized to avoid 

nonspecific recognition of excess proteins. 

The antibodies used for immunoprecipitation 

(Basic Protocol 2) should not interfere with 

the enzymatic activity of the enzymes tested. 

In addition, it is necessary to avoid nonspecific 

precipitation of contaminant kinases. However, 

sometimes the washings may not prevent 

coimmunoprecipitation of protein kinases 

other than those desired, and these might interfere 

with the phosphorylation reaction. This 

difficulty can be overcome by using a specific 

substrate or direct assay methods (i.e., in-gel 

kinase assay, Basic Protocol 3). 

For accurate comparison of the amounts 

of phosphoproteins, detection should be performed 

in the linear range of the detection 

system. Thus, the amount of protein loaded 

on the gel, the concentration of primary and 

secondary antibodies, and the time of ECL exposure 

should be optimized in order to reach 

linearity. Alternatively, a standard curve with 

the proteins of interest can be made and serial 

dilutions of the cellular extracts of each 

treatment can be loaded onto the SDS-PAGE 

gel. The blot detection system, e.g., 

ECL- 125 I-, AP-, or biotin-conjugated antibodies, 

should be chosen carefully. Usually, ECL 

has the narrowest linear range of these systems, 

whereas 125 I-antibodies have a relatively 

broad range. The AP detection system, which 

has moderate linear range, is usually used for 

the types of experiments described here, because 

it is a reliable method that does not necessitate 

the use of radioactive materials. 

Other parameters that should be considered 

for accurate comparison of protein kinases are 

as follows. 



14.3.27 




14.3.28 

1. Starvation of the cells before activation, 

which may interfere with the activation of the 

desired protein kinase or cause activation of 

some stress-activated protein kinases. 

2. The optimal length of stimulation may 

vary from cell type to cell type and from one 

protein kinase to the other; thus, appropriate 

time points for each kinase should be determined. 

3. For accurate comparison of protein kinase 

activities, detection should be performed 

in the linear range of the phosphorylation reaction. 

Thus, the amount of protein used for 

immunoprecipitation, the concentration of antibodies, 

the length of the phosphorylation reaction, 

and the exposure to X-ray film or to the 

phosphor imager should be optimized in order 

to reach linearity. If necessary, a standard 

curve with the protein kinases of interest can 

be made, and serial dilutions of the cytosolic 

extracts or a time course of the phosphorylation 

can be used to ensure one is working in a 

linear range. 

Anticipated Results 

With immunoblotting, the desired phosphoprotein 

should be selectively detected by both 

anti-phospho and anti-general antibodies. For 

example, when anti-active ERK antibodies are 

used (Basic Protocol 1), two faint bands at 

molecular weights of 42 and 44 kDa should be 

detected in the basal, nonstimulated fractions 

of the cells (Fig. 14.3.2). These two bands represent 

the small amount of active ERK (ERK1 

and ERK2) present in resting cells. Upon addition 

of EGF to these cells, the intensity of 

staining of the same bands should increase 

with time, peak at 30 min after stimulation, 

and decline thereafter. Usually a third band at 

46 kDa (ERK1b) appears at the longer time 

points of EGF stimulation. The kinetics of detection 

of the phosphoprotein result from the 

transient activation of ERK by EGF in these 

cells. Staining the same blots with anti-general 

ERK antibodies should result in equal staining 

of the 42-, 44-, and 46-kDa bands in all lanes 

of the blot. 

In Basic Protocol 2, extracts from nonstimulated 

Rat-1 cells should yield very little MBP 

phosphorylation (Fig. 14.3.2), which represents 

the activity of both ERK1 and ERK2 

in these cells. Upon addition of EGF to the 

cells, the amount of phosphate incorporated 

should increase with time, peaking at 30 min 

after stimulation and declining thereafter. The 

kinetics of phosphorylation represent the transient 

activation of ERK by EGF in these cells. 

When the in-gel assay is used (Basic Protocol 

3), both ERK1 and ERK2 should be detected 

at molecular weights of 44 and 42 kDa, and the 

kinetics of activation of both of them should be 

similar to those of the MBP phosphorylation 

obtained with Basic Protocol 1. Typical results 

of Basic Protocol 3 are shown in Figure 14.3.2. 

Time Considerations 

After cell harvesting, Basic Protocol 1 requires 

extraction (0.5 hr), determination of 

protein concentration (0.5 hr), SDS-PAGE 

(2.5 hr), and immunoblot analysis (6 to 8 hr). 

Because this procedure may take more than 

1 working day, it can be stopped after boiling 

the samples in sample buffer, or the transfer 

onto the nitrocellulose membrane can be 

performed overnight at 40 to 50 mA (instead 

of 200 to 300 mA). Immunoprecipitation and 

washes (Basic Protocol 2) add about 4 to 5 hr 

to Basic Protocol 1. 

After cell harvesting, Basic Protocol 2 requires 

extraction (0.5 hr), immunoprecipitation 

and washings (3 to 4 hr), the phosphorylation 

reaction (0.5 to 1.0 hour), SDS-PAGE 

(2.5 hr), and processing of the gel (6 to 16 hr). 

Because this procedure usually takes more 

than 1 working day, it can be interrupted after 

boiling the samples in sample buffer or at any 

time during the destaining period. 

The in-gel kinase assay (Basic Protocol 3) 

may take 2 to 3 days. 

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Contributed by Yoav Shaul and 

Rony Seger 

Department of Biological Regulation, 

The Weizmann Institute of Science 

Rehovot, Israel 


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Phosphoamino Acid Analysis 

It is often valuable to identify the phosphorylated residue in a protein. In the case of 

proteins phosphorylated at serine, threonine, or tyrosine, this is readily accomplished by 

partial acid hydrolysis in HCl followed by two-dimensional thin-layer electrophoresis of 

the labeled phosphoamino acid (see Basic Protocol). Phosphothreonine and phosphotyrosine 

are more stable to hydrolysis in alkali than are RNA and phosphoserine. Therefore, 

mild alkaline hydrolysis of protein samples can be used to enhance the detection of 

phosphothreonine and phosphotyrosine (see Alternate Protocol). 

Although this procedure can be carried out with a protein eluted from a preparative gel 

and concentrated by trichloroacetic acid (TCA) or acetone precipitation, it is most easily 

accomplished by transfer of the protein of interest to a PVDF membrane. This technique 

is obviously not ideal if the protein being studied does not transfer efficiently. 

NOTE: Wear gloves and use blunt-end forceps to handle membranes. 

CAUTION: When working with radioactivity, take appropriate precautions to avoid 

contamination of the experimenter and the surroundings. Carry out the experiment and 

dispose of wastes in an appropriately designated area, following the guidelines provided 

by the local radiation safety department (also see APPENDIX 1D). 

ACID HYDROLYSIS AND TWO-DIMENSIONAL ELECTROPHORETIC 

ANALYSIS OF PHOSPHOAMINO ACIDS 

The protein to be acid hydrolyzed is transferred to a PVDF membrane using the same 

technique used for immunoblotting (UNIT 6.2) or for microsequencing. It is valuable, but 

not absolutely essential, to keep the filter wet following transfer. Following acid hydrolysis, 

phosphoamino acids are separated by two-dimensional thin-layer electrophoresis. 

Because electrophoresis equipment differs considerably in design, the details of the 

assembly and placement of the plate are not discussed here. It is assumed that a suitable 

apparatus is available for use by an experienced operator. Electrophoresis conditions are 

described for using the HTLE 7000 (CBS Scientific). They are almost certainly not correct 

for other equipment and will need to be altered according to the equipment manufacturer’s 

directions. 

Materials 

32P-labeled phosphoprotein (UNIT 14.4) 

India ink solution: 1 µl/ml India ink in TBS (UNIT 14.4)/0.02% (v/v) Tween 20, 

pH 6.5 (prepare fresh or store indefinitely at room temperature); or radioactive 

or phosphorescent alignment markers 

6 M HCl 

Phosphoamino acid standards mixture (see recipe) 

pH 1.9 electrophoresis buffer (see recipe) 

pH 3.5 electrophoresis buffer (see recipe) 

0.25% (w/v) ninhydrin in acetone in a freon (aerosol, gas-driven) atomizer/sprayer 

PVDF membrane (Immobilon-P, Millipore) 

110° oven 

Screw-cap microcentrifuge tubes 

20 cm × 20 cm × 100 µm glass-backed cellulose thin-layer chromatography 

plate (EM Sciences) 

Large blotter: two 25 × 25–cm layers of Whatman 3MM paper sewn together at 

the edges, with four 2-cm holes that align with the origins on the TLC plate 

Contributed by Bartholomew M. Sefton 


Copyright © 1999 by John Wiley & Sons, Inc. 

UNIT 14.5 

BASIC 

PROTOCOL 





14.5.1 

Supplement 3

Phosphoamino 

Acid Analysis 

14.5.2 

Glass tray or plastic box 

Whatman 3MM paper 

Thin-layer electrophoresis apparatus (e.g., HTLE 7000, CBS Scientific) 

Fan 

Small blotters: 4 × 25–cm, 5 × 25–cm, and 10 × 25–cm pieces of Whatman 

3MM paper 

50° to 80°C drying oven 

Sheets of transparency film for overhead projector 

Additional reagents and equipment for SDS-PAGE (UNIT 6.1), immunoblotting (UNIT 

6.2), and autoradiography (UNIT 6.3) 

Prepare sample 

1. Run radiolabeled phosphoprotein on a preparative SDS-polyacrylamide gel (UNIT 6.1). 

It is difficult to obtain good results with 0.5 ml water. 

Place the piece of filter paper in a screw-cap microcentrifuge tube. 

Keep the excised piece as small as possible. 

Hydrolyze sample 

5. Add enough 6 M HCl to submerge the piece of filter. Screw the cap on the tube tightly 

and incubate 60 min in 110°C oven. 

6. Let cool. Microcentrifuge 2 min at maximum speed, room temperature. Transfer the 

liquid hydrolysate to a fresh microcentrifuge tube and dry with a Speedvac evaporator. 

Drying takes ∼2 hr. Simultaneous drying of the hydrolysate and deblocked oligonucleotides 

in NH 4 OH must be avoided, as this will generate a cloud of ammonium chloride that will 

collect in the centrifuge tube and render the hydrolysate unsuitable for thin-layer electrophoresis. 

7. Dissolve the sample in 6 to 10 µl water by vortexing vigorously. Microcentrifuge 5 

min at maximum speed. 

Prepare plate for first-dimension electrophoresis 

8. Spot 25% to 50% of the sample, in 0.25- to 0.50-µl aliquots, on one origin of a 20 

cm × 20 cm × 100 µm glass-backed cellulose thin-layer chromatography plate (see 

Fig. 14.5.1 for arrangement of samples). Between each application, dry the sample 

spot with compressed air delivered through a Pasteur pipet plugged with cotton. 

Use long, thin plastic micropipet loading tips for loading, and do not let the tip touch the 

plate. 

Four samples can be analyzed simultaneously. The complete hydrolysate can be spotted on 

a single origin, but some streaking in the first dimension may be observed due to 

overloading. This problem can be avoided by using a fraction of the sample. 


A 

C 

8 cm 3 cm 

plate with samples 

applied at 4 origins (+) 

blotter applied to plate 

in order to wet it 

8 cm 

3 cm 

9. Spot 1 µl nonradioactive phosphoamino acid standards mixture (containing phosphoserine, 

phosphothreonine, and phosphotyrosine) on top of each sample in 0.25to 

0.50-µl aliquots as above. 

10. Wet the large blotter (with four holes) by submerging it in pH 1.9 electrophoresis 

buffer in a large glass tray or plastic box. Briefly allow the excess buffer to drain off. 

Lower the wet blotter onto the prespotted plate with the origins on the plate in the 

centers of the four holes in the blotter (Fig. 14.5.1). Press on the blotter gently to 

achieve even wetting of the cellulose and concentration of the samples. When the 

plate is uniformly wet, remove the blotter. 

B 

D 

25 cm 

first-dimension blotter 

for electrophoresis at pH 1.9 

phosphoamino acids 

results of first-dimension 

electrophoresis at pH 1.9 

25 cm 

Figure 14.5.1 First-dimension electrophoretic separation of phosphoamino acids at pH 1.9. (A) 

Positions of the four origins on a single 20 × 20–cm plate; (B) blotter used for wetting the plate with 

pH 1.9 electrophoresis buffer; (C) placement of the blotter on the plate (underneath; indicated by 

dashed outline); and (D) orientation of the plate between the + and − electrodes with the positions 

of the phosphoamino acids after electrophoresis. 

The blotter should be quite damp but not sopping wet. Excess buffer can be wicked off onto 

filter paper. 

Areas of the plate that are too dry can be seen through the blotter and will appear to be 

whiter than the rest of the plate. If this happens, dab the blotter with a Kimwipe wetted 

with pH 1.9 electrophoresis buffer. If there are puddles of buffer on the plate, let them dry 

before carrying out electrophoresis. 





14.5.3 


Phosphoamino 

Acid Analysis 

14.5.4 

A 

C 

25 cm 

second-dimension blotters 

for electrophoresis at pH 3.5 

90 o counterclockwise 

rotation of plate 

10 cm 

5 cm 

4 cm 

11. Place the thin-layer plate in the electrophoresis apparatus and overlap 0.5 cm of the 

right and left sides of the plate with wicks made of Whatman 3MM paper. If the 

apparatus has an air bag, be sure to inflate it. Close the cover and start electrophoresis. 

With an HTLE 7000, double-thickness Whatman 3MM wicks, and a plate with four 

samples, electrophorese 20 min at 1.5 kV. 

B 

D 

blotters applied to plate 

in order to wet it 

phosphamino acids 

results of second-dimension 

electrophoresis at pH 3.5 showing 

ninhydrin-stained standards 

Figure 14.5.2 Second-dimension electrophoretic separation of phosphoamino acids at pH 3.5. (A) 

The three pieces of Whatman 3MM paper used for wetting the plate with pH 3.5 electrophoresis buffer; 

(B) proper placement of the blotters on the plate (underneath; indicated by dashed outline); (C) 

reorientation of the plate for electrophoresis in the second dimension; and (D) orientation of the plate 

between the + and - electrodes with the position of the phosphoamino acids after electrophoresis. 

For the HTLE 7000 apparatus, use folded-over Whatman 3MM wicks that are 20 cm wide 

(the same as the plate) and not overly wet. Overly wet wicks will flood the plate and cause 

sample diffusion. 

For other electrophoresis apparatuses the appropriate duration of electrophoresis can be 

determined empirically by examining the rate of migration of the phosphoamino acid 

standards. 


pH 3.5 

pH 1.9 

12. Following electrophoresis, remove the plate and quickly air dry with a fan without 

heating. 

It takes ∼20 min to dry the plate. 

phosphoserine 

phosphothreonine 

phosphotyrosine 

phosphopeptides generated 

by partial hydrolysis 

origin 

Figure 14.5.3 Hypothetical autoradiogram of a two-dimensional separation. Four samples of 

acid-hydrolyzed, 32 P-labeled proteins are applied at the origins, one in each of the four quandrants. 

This diagram shows the origins, the directions of electrophoresis, the positions of phosphoserine, 

phosphothreonine, and phosphotyrosine, the position of P i, and the position of partially hydrolyzed 

fragments of the proteins for the upper right-hand sample. Every protein generates different partial 

hydrolysis peptide fragments. 

Perform second-dimension electrophoresis 

13. Wet the small blotters in pH 3.5 electrophoresis buffer and use them to wet the plate using 

the method described in step 10 to achieve even wetting without puddling (Fig. 14.5.2). 

After electrophoresis at pH 1.9, phosphoamino acids are present as a streak extending from 

the origin towards the + electrode. Blotters are not applied directly over the phosphoamino 

acids to prevent sample blurring or smearing. 

14. Remove the blotters, rotate the plate 90° counterclockwise, and electrophorese 16 

min at 1.3 kV in pH 3.5 electrophoresis buffer if using the HTLE 7000 apparatus. 

15. At the end of the electrophoresis run, remove the plate and dry 20 to 30 min in an 

oven at 50° to 80°C. When dry, spray with 0.25% ninhydrin in acetone, then reheat 

in the oven 5 to 10 min to visualize the phosphoamino acid standards. 

16. Place radioactive or phosphorescent alignment marks on the plate and autoradiograph 

with an intensifying screen overnight to 10 days at −70°C. 

17. Following autoradiography, trace the alignment markers and the stained phosphoamino 

acid markers onto a transparent sheet used for overhead projectors. Save this template. 

Align the film with the plate and identify radioactive phosphoamino acids (Fig. 14.5.3). 

P i 

Use of fluorography or autoradiography to detect the labeled phosphoamino acids is 

preferable to use of a phosphorimager. The image on film is precisely the same size as the 

thin-layer plate, which allows the transparent film to be overlaid on the plate for an 

unambiguous spot identification. A phosphorimager can subsequently be used for quantification. 





14.5.5 


ALTERNATE 

PROTOCOL 

Phosphoamino 

Acid Analysis 

14.5.6 

ALKALI TREATMENT TO ENHANCE DETECTION OF TYR- AND 

THR-PHOSPHORYLATED PROTEINS BLOTTED ONTO FILTERS 

Phosphothreonine and phosphotyrosine are much more stable to hydrolysis in alkali than 

RNA or phosphoserine. Detection of proteins containing phosphothreonine and phosphotyrosine 

in impure samples containing 32P-labeled RNA and serine-phosphorylated 

proteins can often be enhanced by mild alkaline hydrolysis of gel-fractionated samples. 

Although this technique was first developed for the treatment of fixed polyacrylamide 

gels, it is much more easily performed with proteins that have been first transferred to a 

PVDF membrane. 

Alkaline hydrolysis does not preclude subsequent phosphoamino acid analysis. A band 

from a blot that has been treated with alkali can be excised and subjected to acid hydrolysis 

as described in the Basic Protocol. 

Additional Materials (also see Basic Protocol) 

1 M KOH 

TN buffer: 10 mM Tris⋅Cl (pH 7.4 at room temperature)/0.15 M NaCl 

1 M Tris⋅Cl, pH 7.0 at room temperature 

Covered plastic container (e.g., Tupperware box) 

55°C oven or water bath 

1. Run radiolabeled sample on a preparative SDS–polyacrylamide gel and transfer 

proteins electrophoretically to a PVDF membrane (see Basic Protocol steps 1 and 2). 

A band containing as few as 10 cpm is detectable under optimal conditions with this 

technique. 

A nylon membrane may be used in place of a PVDF membrane, but in that case, the bands 

cannot subsequently be analyzed by acid hydrolysis, as nylon membrane will dissolve in 6 

M HCl. 

2. Wash membrane thoroughly with water: three 2-min incubations in 1 liter water are 

sufficient. 

These washes remove buffer and detergent. 

3. Incubate membrane 120 min at 55°C in an oven or water bath in sufficient 1 M KOH 

to cover the filter in a covered Tupperware container. 

4. Discard KOH. Wash membrane and neutralize remaining KOH by rinsing once for 5 

min in 500 ml TN buffer, once for 5 min in 500 ml of 1 M Tris⋅Cl (pH 7.0), and twice 

for 5 min in 500 ml water. Wrap the membrane in plastic wrap and autoradiograph 

(UNIT 6.3) overnight with flashed film and an intensifying screen at −70°C. 

Identification of the band of interest is most easily accomplished by coelectrophoresis of a 

radioactive marker protein of known identity. 


Use deionized or distilled water in all recipes and protocol steps. For common stock solutions, see 

APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX. 

pH 1.9 electrophoresis buffer 

50 ml 88% formic acid (0.58 M final) 

156 ml glacial acetic acid (1.36 M final) 

1794 ml H 2O 

Store indefinitely in a sealed bottle at room temperature 


pH 3.5 electrophoresis buffer 

100 ml glacial acetic acid (0.87 M final) 

10 ml pyridine [0.5% (v/v) final] 

10 ml 100 mM EDTA (0.5 mM final) 

1880 ml H2O Store indefinitely in a sealed bottle at room temperature 

Phosphoamino acid standards mixture 

Prepare a solution of phosphoserine, phosphothreonine, and phosphotyrosine 

(Sigma) in water at a final concentration of 0.3 mg/ml each. Store in 1-ml aliquots 

indefinitely at −20°C. 

COMMENTARY 


Phosphoamino acid analysis by the two-dimensional 

electrophoretic technique described 

in the basic protocol was first carried out with 

proteins isolated by elution from unfixed SDSpolyacrylamide 

gels (Hunter and Sefton, 

1980). However, this technique is laborious, 

especially if it involves grinding up pieces of 

high-percentage acrylamide gels, and the yields 

can be disappointing. Additionally, because the 

eluted protein must be precipitated in the presence 

of a carrier protein, spotting the whole 

sample on a single origin usually yields a badly 

smeared pattern. The grind-and-elute technique 

is, however, advantageous with proteins 

that are very refractory to electrophoretic transfer 

to PVDF membranes. 

The alkaline treatment of protein described 

in the alternate protocol was first developed by 

Jon Cooper and Tony Hunter, who treated fixed 

gels with KOH (Cooper and Hunter, 1981). The 

original technique is tricky because the gel 

becomes extremely sticky during incubation 

with KOH and swells. Additionally, the manipulations 

needed to recover proteins from the 

gel following treatment are very involved because 

the proteins are contaminated with products 

of the hydrolysis of polyacrylamide. 

Critical Parameters and 

Troubleshooting 

It is essential to use PVDF membranes to 

immobilize proteins for acid hydrolysis rather 

than nylon or nitrocellulose membranes, both 

of which dissolve in 6 M HCl. Proteins immobilized 

on either PVDF or nylon membranes 

may be subjected to alkaline hydrolysis with 

KOH (Contor et al., 1987), but nitrocellulose 

membranes are not suitable. Proteins immobilized 

on nylon cannot subsequently be analyzed 

by acid hydrolysis because nylon is dissolved 

by 6 M HCl. 

Two-dimensional thin-layer electrophoresis 

is required for unambiguous identification of 

phosphorylated residues, as some spots after 

one-dimensional electrophoresis do not represent 

pure species. For example, uridine monophosphate, 

which is generated during acid hydrolysis 

of RNA (a frequent contaminant of 

phosphoproteins), comigrates with phosphotyrosine 

during one-dimensional electrophoresis 

at pH 3.5. 

Streaking of the sample in the first dimension 

is a symptom of overloading, either with 

the phosphoprotein itself or with contaminants 

in the sample. This problem can be corrected 

by loading less sample. Streaking in the second 

dimension is usually the result of problems with 

wetting or running the plate and cannot be 

corrected by loading less sample. 

Some batches of blotting paper contain calcium, 

which interferes with electrophoresis of 

phosphoamino acids (probably by precipitating 

them). In the author’s experience Whatman 

3MM paper is quite reliable; other blotting 

papers are probably suitable as well. Inclusion 

of EDTA in the pH 3.5 buffer alleviates this 

problem. 

This unit calls for glass-backed cellulose 

thin-layer plates rather than the plastic-backed 

variety, which are lighter and less expensive. 

This is because plastic-backed plates can under 

some circumstances cause sample streaking. 

They are, however, probably satisfactory for 

most experiments. If use of plastic-backed 

plates results in streaking, try glass-backed 

plates to see if that corrects the problem. 


To detect a phosphoamino acid by autoradiography, 

a minimum of 10 cpm must be 

spotted and the plate exposed for a week with 

flashed film and an intensifying screen. Only 

15% to 20% of the radioactivity in a phospho- 





14.5.7 


Phosphoamino 

Acid Analysis 

14.5.8 

protein is recovered as phosphoamino acids. 

The majority is present as 32 Pi, which is released 

by dephosphorylation of phosphoamino acids, 

with the remainder being peptide products resulting 

from partial acid hydrolysis. As a result, 

the thin-layer plates will contain a number of 

radioactive spots that are not phosphoamino 

acids (see Fig. 14.5.3). Partial hydrolysis products 

remain near the origin during electrophoresis 

at pH 1.9, but exhibit some mobility at pH 

3.5 (see Fig. 14.5.3). After two-dimensional 

electrophoresis, they are found above the origin 

and below the phosphoamino acids. 32 Pi has a 

high mobility at both pH 1.9 and pH 3.5 and is 

found in the upper left-hand corner of each 

quadrant of the plate (see Fig. 14.5.3). Because 

of these additional radioactive spots, it is essential 

to localize internal phosphoamino acid 

standards by staining with ninhydrin. 


After the preparative gel has been run and 

the protein transferred to the membrane, isolation 

of the membrane fragment containing the 

protein, followed by acid hydrolysis, takes

Determination of Akt/PKB Signaling 

Akt—also known as protein kinase B (PKB)—is a central regulator of cell survival 

(Franke et al., 1997; Hemmings, 1997; Marte and Downward, 1997), and its activity is 

often used to assess the apoptotic effect of different experimental conditions. This Ser/Thr 

protein kinase is activated downstream of phosphoinositide 3-kinase (PI3K) in response 

to a wide variety of growth factors (Fig. 14.6.1). The activated form of the kinase targets 

a specific set of effector proteins involved in cell-survival signaling. The activation 

involves membrane translocation and dual phosphorylation on threonine at position 308 

and serine at position 473. Because the activation of Akt depends on this phosphorylation, 

it is possible to assess Akt activity not only with a kinase assay but also by determining 

its level of phosphorylation. 

This unit provides a protocol for assaying the phosphorylation and the dynamics of 

dephosphorylation of Akt/PKB in cultured cells (see Basic Protocol). A protocol is also 

provided for assaying membrane translocation in response to Akt activation (see Support 

Protocol). 

DETERMINATION OF Akt/PKB SIGNALING IN CULTURED CELLS 

This protocol can be applied to nearly all cultured cell lines with few or no modifications. 

In this particular example, the chosen cell system involves cells expressing or deficient 

in the β1 integrin because of the links between integrin signaling, Akt activation, and cell 

survival. This protocol describes PDGF stimulation of GD25 β1 integrin–null cells 

(Fassler et al., 1995) expressing wild-type β1 integrin or its mutant variant W/A, which 

is deficient in Akt signaling (Pankov et al., 2003). Specific treatments and time-course 

sampling of cultured cells are described; these allow determination of the phosphorylation 

level of Akt, its sensitivity to growth factor stimulation, and the mode of its inactivation 

by dephosphorylation. All these results can be obtained in a single nonradioactive 

experiment by using standard techniques such as immunoblotting (UNIT 6.2) and immunodetection 

with phosphospecific antibodies (UNIT 14.2). Cell cultures are serum-starved, 

stimulated with growth factor, and maintained for 2 hr after the stimulation for assessing 

the dynamics of Akt dephosphorylation. Samples are taken after each treatment, and Akt 

activity is determined with phosphospecific antibodies. 

Materials 

GD25 cells obtained from R. Fassler, Max Planck Institute of Biochemistry, 

Martinsried, Germany, and expressing wild-type β1 integrin (β1 cells) or 

integrin mutant W/A (mutant cells) 

Dulbecco’s modified Eagle medium supplemented with 10% fetal bovine serum 

(DMEM/10% FBS; APPENDIX 2A) 

Starvation medium: DMEM containing 1% bovine serum albumin (DMEM/1% 

BSA) 

0.5 µM okadaic acid (OA; 1 mM stock solution in DMSO, APPENDIX 1B) in 

DMEM/1% BSA 

Platelet-derived growth factor-BB (PDGF-BB), freshly prepared at 10 µg/ml in 10 

mM acetic acid 

AG 1433 (e.g., Calbiochem) PDGF kinase inhibitor, 10 mM stock solution in 

dimethyl sulfoxide (DMSO) 

PBS (APPENDIX 2A), ice cold 

Modified radioimmunoprecipitation assay (mRIPA) buffer (see recipe), ice cold 

2× SDS sample buffer (APPENDIX 2A) 

10% separating gels with 4% stacking gels (UNIT 6.1) 

Contributed by Roumen Pankov 


Copyright © 2003 by John Wiley & Sons, Inc. 

UNIT 14.6 

BASIC 

PROTOCOL 





14.6.1 

Supplement 22

Determination of 

Akt/PKB 

Signaling 

14.6.2 

PI3K 

PH 

T308 

Akt/PKB 

Ptdlns(3,4,5)P3 PH 

S473 

PDK1 

T308- P 

Akt/PKB 

phosphorylation 

(activation) 

PP2A 

dephosphorylation 

(inactivation) 

OA 

PDK2 

S473- P 

membrane 

Akt/PKB 

Pre-stained protein standards (e.g., Novex) 

Transfer buffer (UNIT 6.2) 

Ponceau S solution (UNIT 6.2) 

Tris-buffered saline with 0.1% Tween 20 (TTBS, APPENDIX 2A) 

Blocking solution: TTBS containing 5% dry nonfat milk (TTBS/milk) 

Primary antibodies: polyclonal anti-phospho Akt (Ser 473) (e.g., Cell Signaling, 

Biosource), monoclonal anti-actin (e.g., Sigma) 

Dry nonfat milk 

Secondary horseradish peroxidase (HRP)–conjugated anti-rabbit or anti-mouse 

antibodies (e.g., Amersham Biosciences) 

Enhanced chemiluminescence (ECL) detection reagent (UNIT 14.2) 

35-mm tissue culture dishes 

Plastic cell scraper (rubber policeman) 

1.5-ml microcentrifuge tubes, prechilled 

Sonicator/ultrasonic processor 

Boiling water bath 

Two nitrocellulose membranes cut to gel size 

PH 

GSK3 

Caspase 9 

Forkhead 

family 

T308- P 

Bad 

S473- P 

p70S6K 

eNOS 

Figure 14.6.1 Activation of Akt/PKB and its effectors. Akt is activated by phosphatidylinositol 

3-kinase (PI3K) products PtdIns(3,4,5)P 3 and PtdIns(3,4,)P 2. These phosphoinositides target 

inactive Akt to the plasma membrane via its pleckstrin homology (PH) domain. At the membrane, 

Akt is phosphorylated on two residues: threonine 308 (T308) by phosphoinositide-dependent kinase 

1 (PDK1) and serine 473 (S473) by an unidentified PDK2. The dual phosphorylation is necessary 

for full activation of Akt. Activated Akt in turn phosphorylates its targets, thus modulating their 

function. Akt effectors include glycogen synthase kinase-3 (GSK3), caspase 9, Forkhead family of 

transcription factors, Bad, p70 ribosomal S6 kinase (p70S6K), IκB kinases (IKKs), and endothelial 

nitric oxide synthase (eNOS). Akt is inactivated through dephosphorylation by protein phosphatase 

2A (PP2A), and this process can be blocked by okadaic acid (OA). 

Supplement 22 Current Protocols in Cell Biology 

IKKs

Four Whatman 3MM filter papers cut to gel size 

SDS-PAGE/transfer apparatus (e.g., Bio-Rad, Novex) 

Constant-voltage/current power supply (e.g., Bio-Rad) 

Flat containers for washing gels and membranes 

Shaker 

Heat-sealable plastic bags 

Heat sealer 

X-ray film (e.g., Hyperfilm; Amersham Biosciences) 

Film cassette for X-ray film 

X-ray film developer 

Additional reagents and equipment for tissue culture (UNIT 1.1), SDS-PAGE (UNIT 6.1), 

and immunoblotting (UNIT 6.2) 

NOTE: All tissue culture incubations should be performed in a humidified 37°C, 10% 

CO 2 incubator. Use pre-warmed cell culture medium for all treatments. 

Treat cells 

1. Plate twelve 35-mm tissue culture dishes each of GD25 cells expressing wild-type 

β1 integrin and mutant cells (1.0 × 10 6 cells/dish) in 2 ml DMEM/10% FBS and allow 

them to attach and spread for 3 to 4 hr. 

2. Wash dishes with 2 ml starvation medium two times, then add 2 ml/dish of the 

starvation medium and serum-starve the cells overnight. 

At this point the dishes should be ∼80% confluent. Depending on the cell type being used, 

adjustment of the initial number of plated cells may be necessary. 

3. Treat half of the dishes (six dishes of β1 cells and six dishes of mutant cells) with 0.5 

µM okadaic acid (OA) (protein phosphatase inhibitor) in DMEM/1% BSA for 15 

min. Label the dishes “OA”. Maintain the same concentration of OA in the medium 

to be used to treat these dishes throughout the entire experiment. To ensure similar 

treatment, dispense 25 ml of pre-warmed DMEM/1% BSA and add 12.5 µl of 1 mM 

okadaic acid stock solution to the 25 ml of medium to be used for the “OA” sets of 

dishes. Add 12.5 µl of DMSO to the other 25 ml medium for the untreated sets. 

After this treatment, four sets of six samples are formed (two for each cell line). These are 

untreated cells of each β1 cells (β1) and W/A mutant cells (mutant), and okadaic acid– 

treated β1 cells (β1/OA) and mutant cells (mutant/OA). 

4. Aspirate the starvation medium and stimulate cells on five dishes of each set with 20 

ng/ml PDGF-BB in 2 ml DMEM/1% BSA for 15 min. Leave four dishes, one from 

each set (β1, β1/OA, W/A, and W/A/OA) without stimulation in fresh DMEM/1% 

BSA and label them as controls. To ensure similar treatment, dispense 40 ml of 

pre-warmed DMEM/1% BSA and add 80 µl of 10 µg/ml stock solution of PDGF-BB, 

mix well, divide into two 20-ml aliquots, and add 10 µl of 1 mM okadaic acid stock 

solution to the 20 ml of medium to be used for the “OA” sets of dishes. Add 10 µl of 

DMSO to the other 20 ml of PDGF-containing medium. 

5. Rinse all dishes one time with DMEM/1% BSA. Put aside the control dishes and one 

of each PDGF-stimulated dishes (β1, β1/OA, W/A, and W/A/OA) for lysis. To the 

remaining four dishes of each set, add 2 ml/dish of the same medium containing 30 

µM AG 1433 PDGF kinase inhibitor (four of the β1 and four of the mutant 

PDGF-stimulated dishes). Treat the “OA” set of dishes (four of the β1/OA and four 

of the W/A/OA PDGF-stimulated dishes) with the same AG 1433–containing medium 

supplemented with 0.5 µM okadaic acid. Incubate the AG 1433–containing 

dishes for 30, 60, 90, and 120 min at 37°C. Specifically, dispense 40 ml of pre-warmed 





14.6.3 



Akt/PKB 


14.6.4 

DMEM/1% BSA and add 120 µl of 10 mM stock solution of AG 1433, mix well, and 

divide into 20-ml aliquots and add 10 µl of 1 mM okadaic acid stock solution to the 

20 ml of medium to be used for the “OA” sets of dishes. Add 10 µl of DMSO to the 

other 20 ml of AG 1433-containing medium. 

AG 1433 is used to eliminate residual receptor kinase activity after the PDGF stimulation 

and to identify the rate of dephosphorylation. 

Prepare cellular lysates 

6. Rinse the dishes that were put aside for lysis (control and PDGF-stimulated β1, 

β1/OA, W/A, and W/A/OA) with 2 ml ice-cold PBS two times. Keep the dishes on 

ice, and use ice-cold buffers to slow down biological processes and prevent protein 

degradation. 

7. Add 100 µl/dish of ice-cold mRIPA buffer plus inhibitors, scrape the cells with a 

plastic scraper or rubber policeman, and transfer the lysate into prechilled and 

prelabeled 1.5-ml microcentrifuge tubes. Incubate the lysates for an additional 10 

min on ice. 

Prepare gel samples 

8. Add 100 µl/sample of 2× SDS sample buffer, sonicate (two 5-sec, 50-W pulses) on 

ice. 

The samples will become quite viscous after the addition of the SDS sample buffer. It is 

necessary to shear the released DNA by sonication to eliminate the viscosity and to allow 

correct loading of the samples on SDS-PAGE gels. 

9. Boil samples in a water bath for 3 min or heat at 95°C for 5 min on a heating block. 

10. Remove the PDGF-stimulated and AG 1433–treated β1, β1/OA, W/A, and W/A/OA 

dishes after 30, 60, 90, and 120 min. 

11. Rinse with 2 ml ice-cold PBS two times and repeat steps 7, 8, and 9 for the samples 

at each timepoint. 

The samples can be stored sealed and frozen at least 1 month at −20°C. 

Separate samples by SDS-PAGE 

12. Cast two 10% separating gels with 4% stacking gels (UNIT 6.1). 

Use 15-well combs so that all 24 samples can be loaded on two gels. 

13. Load 20 µl of each sample/gel lane and a separate sample containing prestained 

protein standards on the gel. 

Load each set of samples starting with the control sample, followed by the PDGF-stimulated 

sample, taken immediately after the stimulation, followed by the samples that have 

been kept in AG 1433–containing medium for 30, 60, 90, and 120 min for each sample set. 

Load the prestained protein standards into the first or the last lane so that the orientation 

of the gel can be identified after it is removed from the apparatus. Fill the empty lanes of 

the gels with 1× SDS sample buffer to prevent distortion of the separation in the adjacent 

lanes. 

14. Run the gels at 150 V until the bromophenol blue dye reaches the bottom of the gel 

(see UNIT 6.1). 


Transfer separated proteins from gel to membrane 

15. When the electrophoresis is complete, remove the gels from the gel plates, cut off 

the stacking gels and incubate the separating portion of the gels in 50 ml transfer 

buffer for 15 min. 

Use gloves to handle the gels and membranes since oils from hands can block the transfer. 

16. Assemble the transfer sandwich consisting of pad, Whatman 3MM filter paper, 

nitrocellulose membrane, equilibrated acrylamide gel, second Whatman 3MM filter 

paper, and second pad (Fig. 6.2.1). 

All pads, filter papers, and nitrocellulose membranes should be handled using gloves and 

prewetted with transfer buffer. The transfer cassette should be assembled underneath the 

transfer buffer to avoid trapping air bubbles. Ensure the orientation of the gel (judged by 

the position of the prestained protein standards) such that the correct order of the samples 

after transfer onto the nitrocellulose membrane is maintained. 

17. Place the transfer sandwich into the electroblotting apparatus filled with transfer 

buffer with the nitrocellulose membrane on the cathode side of the gel. Connect the 

apparatus to the power supply and transfer proteins for 1 hr at 100V (constant voltage) 

with cooling (UNIT 6.2). 

Transfer time depends on the size of the proteins, acrylamide percentage, and the thickness 

of the gel. The completeness of the transfer can be easily judged by the transfer of the 

prestained protein standards. 

18. At the end of the transfer, turn off the power supply and disassemble the apparatus 

and transfer cassette. Remove the nitrocellulose membranes and stain with 50 ml 

Ponceau S solution in a flat container for 5 min. Destain the membranes by rinsing 

several times with distilled water. 

Two membranes can be incubated in the same container by orienting them back to back. 

Staining with Ponceau S does not interfere with the subsequent antibody reactions and 

provides a good estimation of the protein loading, separation, and quality of the transfer. 

The Ponceau S solution can be reused several times. 

Probe the membranes with antibodies 

19. Rinse the membranes once with TTBS and incubate in 50 ml blocking solution for 

30 min at room temperature with gentle shaking. 

Milk proteins in the blocking buffer are used to saturate free protein-binding sites and to 

prevent the nonspecific binding of the antibody. Do not incubate the membrane longer than 

1 hr in this buffer since blocking buffer also has a slight stripping effect and may cause 

detachment of the transferred sample proteins. 

20. Dilute primary anti-phospho Akt antibody according to the supplier’s instructions in 

10 ml TTBS containing 3% dry nonfat milk. Place the membranes in heat-sealable 

plastic bags, add diluted antibody, and seal the bag with a heat sealer. Incubate the 

membranes overnight at 4°C with gentle shaking. 

Two membranes can be incubated in the same bag by orienting them back to back. Remove 

all air bubbles from the bag before sealing it. 

21. Remove the membranes from the bag and wash them three times, 15 min each with 

50 ml TTBS in flat container with vigorous shaking. 

Do not allow the membranes to dry out after the incubation with primary antibody. 





14.6.5 


SUPPORT 

PROTOCOL 


Akt/PKB 


14.6.6 

22. Dilute the secondary antibody in 10 ml TTBS, 3% dry nonfat milk according to the 

supplier’s instructions. Place the membranes in a new heat-sealable bag, add diluted 

antibody and seal the bag. Incubate for 30 to 45 min at room temperature with gentle 

shaking. 

Either HRP- or AP-conjugated secondary antibodies can be used. HRP-conjugated secondary 

antibodies can be combined with the high-sensitivity ECL detection system. This 

system allows detection of signals from weak antibodies although attention should be paid 

if accurate quantification of the signal is necessary (see Commentary). 

23. Repeat step 21. 

24. Use the ECL immunodetection protocol (UNIT 6.2) to detect phosphorylated Akt. 

Incubate the membranes with ECL solution for 1 min, dry the excess fluid by touching 

the edge of the membrane to a piece of filter paper, wrap the membranes in plastic 

wrap, and expose to X-ray film for 1 min. 

25. Incubate the membranes in 50 ml blocking buffer for 15 min at room temperature 

with shaking. 

26. Repeat steps 20 to 24 with new primary antibody (e.g., anti-actin). 

This second reaction is used as an internal control for loading. 

If both primary antibodies are generated in the same species (e.g., both are mouse or both 

are rabbit) and the molecular weights of the antigens are different (e.g., 60 kDa for Akt and 

45 kDa for actin) the membranes can be incubated with a mixture of the primary antibodies, 

and the two signals can be detected simultaneously on the same X-ray film. 

DETERMINATION OF Akt/PKB TRANSLOCATION TO THE PLASMA 

MEMBRANE 

An important step in Akt activation involves its targeting to the plasma membrane. This 

translocation is dependent on the pleckstrin homology (PH) domain localized at the 

N-terminus of the Akt molecule and the presence of phosphatidylinositol 3-kinase 

products PtdIns(3,4,5)P3 and PtdIns(3,4,)P2. This protocol describes the preparation of membrane and cytosolic fractions from starved 

and PDGF-stimulated GD25 β1 integrin-null cells expressing wild-type β1 integrin or its 

mutant variant W/A that is deficient in Akt signaling. The distribution of Akt is determined 

in each fraction by immunoblotting and immunodetection with antibodies. The same 

protocol can be applied to almost all cultured cell lines with little or no modifications. 

Additional Materials (also see Basic Protocol) 

Cytosolic buffer (see recipe) 

Membrane buffer (see recipe) 

Dry ice 

60-mm dishes 

Treat cells 

1. Plate two 60-mm dishes each of β1 and mutant cells (2.5 × 10 6 cells/dish) in 

DMEM/10% FBS and allow them to attach and spread for 3 to 4 hr. 

2. Wash dishes with 5 ml DMEM/1% BSA (starvation medium) two times, then add 5 

ml/dish of DMEM/1% BSA and serum-starve the cells overnight. 

At this point the dishes should be ∼80% confluent. Depending on the cell type being used, 

adjustment of the initial number of plated cells may be necessary. 


3. Aspirate the starvation medium and stimulate one dish from each cell line with 20 

ng/ml PDGF-BB in 5 ml DMEM/1% BSA for 15 min. Leave one β1 and one mutant 

dish without stimulation in fresh DMEM/1% BSA and label them as controls. To 

ensure similar treatment, dispense 10 ml of pre-warmed DMEM/1% BSA and add 

20 µl of 10 µg/ml stock solution of PDGF-BB, mix well, and add to the dishes to be 

stimulated. 

Prepare cytosol fractions 

4. Rinse the control and PDGF-stimulated dishes with 5 ml cytosolic buffer two times. 

Keep the dishes on ice and use ice-cold buffers to slow down the biological processes 

and prevent protein degradation. 

5. Add 350 µl/dish of ice-cold cytosolic buffer, scrape the cells with a plastic scraper 

or rubber policeman, and transfer the lysate into prelabeled 1.5-ml microcentrifuge 

tubes. 

6. Place the tubes on dry ice until frozen. Thaw the cells in a 37°C water bath. Repeat 

freeze-thaw cycle two additional times. 

Freeze-thaw cycles will break plasma membranes and liberate most of the cytosolic 

proteins. 

7. Centrifuge 15 min at 19,000 × g, 4°C. Transfer the supernatant into prelabeled 1.5-ml 

microcentrifuge tubes, mix with equal volume of 2× SDS sample buffer, and leave 

on ice. 

This supernatant represents the cytosolic fraction. 

Prepare membrane fraction 

8. Resuspend the pellet in fresh 350 µl cytosolic buffer, centrifuge as in step 7, and 

discard the supernatant. 

This washing step clears most of the remaining cytosolic proteins from the pellet. 

9. Suspend the pellet in 50 µl membrane buffer and centrifuge 15 min at 10,000 × g, 

4°C. 

This step solubilizes cellular membranes. Pipet the pellet up and down through a micropipet 

tip several times, but avoid excessive foaming. 

10. Transfer the supernatant into prelabeled 1.5-ml microcentrifuge tubes and mix with 

an equal volume of 2× SDS sample buffer. 

Analyze fractions 

11. Boil all the samples (supernatants from steps 7 and 10) in a boiling water bath for 3 

min or heat at 95°C for 5 min on a heating block. 

The samples can be stored sealed and frozen at least 1 month at −20°C. 

12. Proceed with SDS-PAGE and immunoblotting (see Basic Protocol 1, steps 12 through 

24). Use anti-Akt antibody to probe the distribution of the total Akt between the 

cytosolic and membrane fractions. 





14.6.7 



Akt/PKB 


14.6.8 


Use deionized or distilled water in all recipes and protocol steps. For common stock solutions, see 


Cytosolic buffer 

20 mM Tris⋅Cl, pH 7.5 (APPENDIX 2A) 

150 mM NaCl 

50 µM leupeptin (APPENDIX 1B), add fresh 

50 µM pepstatin (APPENDIX 1B), add fresh 

1 mM PMSF (APPENDIX 1B), add fresh 

1 mM sodium vanadate (APPENDIX 1B), add fresh 

50 mM NaF, add fresh 

Store up to 3 months at 4°C 

The cystolic buffer is derived from Kobayashi et al. (2001). 

Membrane buffer 

20 mM Tris⋅Cl, pH 7.5 (APPENDIX 2A) 

150 mM NaCl 








The membrane buffer is derived from Kobayashi et al. (2001). 

Modified radioimmunoprecipitation assay (mRIPA) buffer 

50 mM HEPES, pH.5 

150 mM NaCl 


0.1% (w/v) SDS (APPENDIX 2A) 

1% (w/v) sodium deoxycholate 


1.5 mM MgCl 2 

1 mM EGTA 







COMMENTARY 


Akt (c-Akt) is the cellular homolog of the 

viral oncoprotein v-Akt. It was first identified 

as a protein kinase with high homology to 

protein kinases A and C and was therefore 

termed PKB (protein kinase B) or RAC (related 

to A and C). Mammals have three closely related 

Akt genes that are expressed differentially 

at both mRNA and protein levels (Datta et al., 

1999; Brazil and Hemmings, 2001). The family 

of Akt proteins contains a central kinase domain 

with specificity for serine or threonine residues 

in substrate proteins and a carboxyl terminus 

containing a hydrophobic motif (HM) and a 

proline-rich domain. The N-terminal region of 

the molecule includes a pleckstrin homology 

(PH) domain, which together with the HM play 

important roles in the Akt activation process 

(Scheid and Woodgett, 2003). The activation 

mechanism involves direct binding of PI3K 


products phosphatidylinositol 3,4,5-trisphosphate 

and phosphatidylinositol 3,4-bisphosphate 

to the PH domain of Akt (Chan et al., 

1999). As a consequence, Akt is localized to the 

plasma membrane where PI3K-generated 3′phosphorylated 

phospholipids reside (Fig. 

14.6.1). This translocation is now known to be 

an important step in Akt activation (Testa and 

Bellacosa, 1997). Myristylated c-Akt, which is 

specifically targeted to the plasma membrane, 

is constitutively active similarly to the oncogenic 

v-Akt, which is permanently targeted to 

the plasma membrane by the viral gag sequence, 

and exhibits constant kinase activity. 

Membrane-localized Akt undergoes conformational 

changes leading to the exposure and 

phosphorylation of two residues—Thr308 in 

the activation loop, proximal to the catalytic 

core, and Ser473 in the HM. This dual phosphorylation 

is necessary for the full activation 

of Akt. The kinase that phosphorylates Thr308 

has been named 3-phosphoinositide-dependent 

kinase 1 (PDK1) because it also requires lipids 

for its activity (Vanhaesebroeck and Alessi, 

2000). Phosphorylation of Akt on Thr308 

causes a charge-induced change in conformation 

allowing substrate binding and an elevated 

rate of catalysis. The second important phosphorylation 

event associated with Akt activation 

occurs at Ser473. The mechanism of this 

phosphorylation is not completely understood 

and may involve autophosphorylation or distinct 

serine kinases like integrin-linked kinase 

(ILK; Persad and Dedhar, 2003). The function 

of phosphorylation at this site is also not fully 

clarified, though the necessity of this modification 

for Akt activity is well documented. For 

example, kinase activity is significantly reduced 

by mutations of this residue, and a similar 

effect is observed after ceramide-promoted 

dephosphorylation at this site. It has been proposed 

that phosphorylation at Ser473 may 

change the properties of the hydrophobic motif, 

shifting it to a docking site for PDK1 (Scheid 

and Woodgett, 2003). After activation, Akt appears 

to detach from the inner leaflet of the 

plasma membrane and to translocate through 

the cytosol to the nucleus (Andjelkovic et al., 

1997; Meier and Hemmings, 1999). 

Cells can inhibit or reverse the activation of 

Akt by several mechanisms (Hill and Hemmings, 

2002). One of them includes the lipid 

phosphatase PTEN (phosphatase and tensin 

homolog deleted on chromosome 10; Yamada 

and Araki, 2001), which decreases the levels of 

phosphatidylinositol 3,4,5-trisphosphate and 

phosphatidylinositol 3,4-bisphosphate within 

cells, thus preventing membrane translocation 

of Akt. Direct inactivation of Akt occurs by 

dephosphorylation of Thr308 and Ser473. This 

process is mediated by the ubiquitous Ser/Thr 

protein phosphatase 2A (PP2A; see Millward 

et al., 1999) and can be blocked by the phosphatase 

inhibitor okadaic acid. A negative regulator 

termed CTMP (carboxy-terminal modulator 

protein), which attenuates the activation 

of Akt at plasma membrane, has also been 

identified (Maira et al., 2001). Thus, the level 

of Akt activity in steady state is the result of an 

equilibrium between activation and inhibition 

events. 

Extensive research has clarified the intracellular 

mechanisms of Akt activation by upstream 

PI3K and PDK1. Less is known about the 

means through which Akt regulates cell growth, 

proliferation, and survival, even though a number 

of downstream Akt substrates have been 

identified (Fig. 14.6.1; Chan et al., 1999; Datta 

et al., 1999; Vanhaesebroeck and Alessi, 2000). 

While the Akt phosphorylation consensus sequence 

is defined (RXRXXS/THydrophobic), 

to date only a small number of Akt substrates 

have been positively or tentatively identified. 

Ongoing studies on Akt signaling and the 

possibility of using this kinase as a drug target 

for cancer, diabetes, and stroke, make probing 

for Akt activity a frequently used task. This can 

be achieved by measuring Akt kinase activity 

directly or by assaying the level of phosphorylation 

as described in this unit. Determination 

of Akt kinase activity follows the general 

scheme applicable for most kinases (e.g., see 

UNIT 14.3) and involves immunoprecipitation of 

the enzyme with antibodies and then performing 

a phosphorylation reaction in vitro. While 

this approach offers a direct and relatively accurate 

determination of Akt activity, the method 

is laborious and often involves use of radioactivity. 

Determination of Akt activation state by 

assaying the level of phosphorylation after specific 

treatments is quicker and employs basic 

non-radioactive laboratory techniques. Moreover, 

the same samples (in some cases the same 

membrane used for immunoblotting with anti- 

Akt antibodies) can be used for determination 

of the activity of upstream Akt regulators and 

the phosphorylation level of downstream Akt 

substrates, making this method more versatile 

than the classical kinase assay. 



Several parameters play critical roles for 

success in determination of Akt activation and 





14.6.9 



Akt/PKB 


14.6.10 

A 

no 

inhibitor 

okadaic 

acid 

B 

– PDGF 

+ PDGF 

β1 cells 

PDGF PDGF 

inhibition. Initial reduction of the phosphorylation 

level of Akt is achieved by a period of 

starvation, which varies depending on the type 

of cells and should be determined experimentally. 

For some cell lines like primary human 

fibroblasts, withdrawal from serum for 4 hr is 

sufficient, while for most immortalized cell 

lines, longer periods of starvation work better. 

The length of serum starvation of the chosen 

cell line needs to be such that it will ensure a 

five-fold or higher increase of Akt phosphorylation 

after stimulation with growth factor. 

mutant cells 

contr 30 60 90 120 contr 30 60 90 120 

β1 cells 

mutant 

cells 

C M C M 

WB total-Akt 

WB 

p-Akt (Ser 473 ) 

actin 

p-Akt (Ser 473 ) 

Figure 14.6.2 Probing of Akt signaling by the methods described in this unit. (A) Determination 

of Akt signaling in cultured cells. GD25 cells expressing either wild-type (β1 cells) or tryptophan 

mutant (mutant cells) integrins were cultured overnight in the absence of serum. Cells were then 

stimulated with 20 ng/ml PDGF-BB for 15 min, the growth factor was washed away, and the cells 

were maintained in medium without serum supplemented with 30 µM PDGF kinase inhibitor AG 

1433. Samples were taken after starvation (contr), immediately after stimulation (PDGF, ↓), and at 

the indicated time points. The experiment was performed in the absence (no inhibitor) or presence 

of 0.5 µM okadaic acid. Lysates from the samples were analyzed by immunoblotting (WB) with 

anti-phospho Ser 473 Akt [p-Akt (Ser 473 )] or anti-actin (actin) antibodies. Actin was used as an 

internal control for loading. (B) Determination of Akt translocation to the plasma membrane. Cells 

were starved overnight and stimulated with 20 ng/ml PDGF (+ PDGF) or left without stimulation 

(–PDGF) and used to prepare cytoplasmic (C) and membrane (M) fractions. Samples from these 

fractions were analyzed by immunoblotting with anti-total Akt (WB total-Akt) antibodies. Increased 

amount of Akt is detected in the membrane fraction of the PDGF stimulated cells. 

actin 

The response to PDGF by different cell lines 

varies significantly. If the increase of Akt phosphorylation 

after stimulation is not sufficient, 

activation with another growth factor, e.g., 

EGF, may be used. If a different growth factor 

is applied, the corresponding growth factor 

receptor (GFR) kinase inhibitor should be 

added to the medium instead of AG 1433 (e.g., 

AG 1478 if EGF is utilized). Addition of a 

growth factor kinase inhibitor is necessary to 

block residual activity of the receptor after 

withdrawal of the growth factor. This treatment 

is critical, since residual GFR activity may 


mask the Akt inactivation pattern by prolonging 

the dephosphorylation time. Experimental 

blocking of dephosphorylation is achieved by 

treatment with okadaic acid. Incubations with 

this potent protein phosphatase inhibitor, especially 

for prolonged times, may cause cell 

rounding and even detachment from the substrate. 

Okadaic acid sensitivity should be determined 

experimentally for each cell type to be 

used. 

Obtaining a high signal-to-noise ratio after 

immunoblotting is essential for the successful 

determination of Akt signaling. This can be 

ensured by: (1) use of specific antibodies that 

recognize the phosphorylated Akt but not its 

unphosphorylated form (now offered by several 

companies like Cell Signaling, Biosource); 

(2) use of freshly added phosphatase and protease 

inhibitors added to the lysis buffers to 

prevent Akt dephosphorylation and degradation 

after cell disruption; (3) loading sufficient 

amount of proteins from the cellular lysate that 

will ensure trouble-free detection of the Akt by 

the antibodies (this is easily achieved by keeping 

the samples for SDS-PAGE concentrated— 

e.g., >5 × 10 5 cells/minigel lane); (4) following 

the proper techniques for SDS-PAGE and immunoblotting 

(see UNITS 6.1 & 6.2). 

Accurate comparison and quantification by 

densitometry of the amounts of phosphorylated 

Akt in different samples can be achieved if the 

detection system is kept in a linear range. The 

enhanced chemiluminescence (ECL) system 

should be optimized to obtain linearity by adjustments 

of the amount of the protein loaded 

on the gel, concentrations of primary and secondary 

antibody, and the X-ray film exposure 

time. If the weakest signal is detectable and the 

strongest signal is still within the linear range 

of the film (e.g., not saturated), then the rest of 

the samples are also in the linear range of the 

system, which can be used for quantification. 


Typical results expected after probing for 

Akt activity (see Basic Protocol) or Akt membrane 

translocation (see Support Protocol) are 

presented in Figure 14.6.2A and B, respectively. 

Comparison between the starved cells 

(contr) and cells after PDGF stimulation 

(PDGF) should demonstrate a several-fold increase 

in the amount of phosphorylated Akt. 

Attenuation of this response after experimental 

manipulations of the cell cultures may indicate 

effects on the upstream pathways leading to Akt 

activation. Evaluation of the amount of phosphorylated 

Akt in the samples taken after PDGF 

stimulation at different time points provides 

information about the rate of Akt dephosphorylation 

(inhibition). While in the samples from 

β1 cells, this decrease is modest, the Akt in the 

samples from mutant cells is rapidly dephosphorylated. 

Such a result indicates activation of 

some of the systems for Akt inhibition (see 

Background Information). Performing the 

same experiment in the presence of a PP2A 

inhibitor, okadaic acid completely reverses this 

effect, indicating that activation of this phosphatase 

is involved in the observed increased 

dephosphorylation rate in W/A mutant cells. If 

OA is ineffective, additional experiments 

should be designed to test the role of other Akt 

inhibitors (see Background Information). Activation 

of Akt is dependent on its membrane 

translocation. A typical increase in membranebound 

Akt after growth factor stimulation is 

presented in Figure 14.6.2B. A modest but 

detectable increase in total Akt in the membrane 

fraction is observed in both cell lines after 

PDGF stimulation. Failure to detect such 

translocation may indicate defects in the function 

of the PH domain of Akt or insufficient 

phosphatidylinositol 3,4,5-trisphosphate and 

phosphatidylinositol 3,4-bisphosphate (see 

Background Information). 


The entire procedure described in the Basic 

Protocol can be completed in 3 days. This 

period includes the time for cell attachment 

after plating (4 hr); starvation (12 hr); PDGF 

stimulation and the necessary incubations up to 

preparation of the SDS-PAGE samples (3 hr); 

SDS-PAGE and electrotransfer (6.5 hr for mini 

gels or 9.5 hr for normal size gels); overnight 

incubation with the primary antibody; and 

completion of the immunoreactions with ECL 

processing (4 hr). Since this procedure takes >1 

day, it is helpful to use one night for starvation 

of the cells and the next night for incubation 

with the primary phosphospecific antibody. 

There are a number of points where the procedure 

can be interrupted: (1) after the preparation 

of the SDS-PAGE samples; (2) after the 

electrotransfer (membranes can be stored wet 

or dry in resealable plastic bags at 4°C); and (3) 

after the completion of the first immunoreaction 

(membranes can be stored wet in resealable 

plastic bags at 4°C). A similar timeframe applies 

for the procedure described in the Support 

Protocol. 





14.6.11 



Akt/PKB 


14.6.12 


Andjelkovic, M., Alessi, D.R., Meier, R., Fernandez, 

A., Lamb, N.J.C., Frech, M., Cron, P., Cohen, 

P., Lucoc, J.M., and Hemmings, B.A. 1997. 

Role of translocation in the activation and function 

of protein kinase B. J. Biol. Chem. 

272:31515-31524. 

Brazil, D.P. and Hemmings, B.A. 2001. Ten years of 

protein kinase B signalling: A hard Akt to follow. 

Trends Biochem. Sci. 26:657-664. 

Chan, T.O., Rittenhouse, S.E., and Tsichlis, P.N. 

1999. AKT/PKB and other D3 phosphoinositide-regulated 

kinases: Kinase activation by 

phosphoinositide-dependent phosphorylation. 

Annu. Rev. Biochem. 68:965-1014. 

Datta, S.R., Brunet, A., and Greenberg, M.E. 1999. 

Cellular survival: A play in three Akts. Genes 

Dev. 13:2905-2927. 

Fassler, R., Pfaff, M., Murphy, J., Noegel, A., Johansson, 

S., Timpl, R., and Albrecht, R. 1995. 

Lack of beta 1 integrin gene in embryonic stem 

cells affects morphology, adhesion, and migration 

but not integration into the inner cell mass 

of blastocysts. J. Cell Biol. 128:979-988. 

Franke, T.F., Kaplan, D.R., and Cantley, L.C. 1997. 

PI3K: Downstream AKT ion blocks apoptosis. 

Cell 88:435-437. 

Hemmings, B.A. 1997. Akt signaling: Linking 

membrane events to life and death decisions. 

Science 275:628-603. 

Hill, M.M. and Hemmings, B.A. 2002. Inhibition of 

protein kinase B/Akt. Implications for cancer 

therapy. Pharmacol. Ther. 93:243-251. 

Kobayashi, S., Shirai, T., Kiyokawa, E., Mochizuki, 

N., Matsuda, M., and Fukui, Y. 2001. Membrane 

recruitment of DOCK180 by binding to 

PtdIns(3,4,5)P3. Biochem. J. 354:73-78. 

Maira, S.M., Galetic, I., Brazil, D.P., Kaech, S., 

Ingley, E., Thelen, M., and Hemmings, B.A. 

2001. Carboxyl-terminal modulator protein 

(CTMP), a negative regulator of PKB/Akt and 

v-Akt at the plasma membrane. Science 

294:374-380. 

Marte, B.M. and Downward, J. 1997. PKB/Akt: 

Connecting phosphoinositide 3-kinase to cell 

survival and beyond. Trends Biochem. Sci. 

22:355-358 

Meier, R. and Hemmings, B.A. 1999. Regulation of 

protein kinase B. J. Recept. Signal Transduct. 

Res. 19:121-128. 

Millward, T.A., Zolnierowicz, S., and Hemmings, 

B.A. 1999. Regulation of protein kinase cascades 

by protein phosphatase 2A. Trends Biochem. 

Sci. 24:186-191. 

Pankov, R., Cukierman, E., Clark, K., Matsumoto, 

K., Hahn, C., Poulin, B., and Yamada, K.M. 

2003. Specific beta 1 integrin site selectively 

regulates Akt/protein kinase B signaling via local 

activation of protein phosphatase 2A. J. Biol. 

Chem. 278:18671-18681. 

Persad, S. and Dedhar, S. 2003. The role of integrinlinked 

kinase (ILK) in cancer progression. Cancer 

Metastasis Rev. 22:375-384. 

Scheid, M.P. and Woodgett, J.R. 2003. Unravelling 

the activation mechanisms of protein kinase 

B/Akt. FEBS Lett. 546:108-112. 

Testa, J.R. and Bellacosa, A. 1997. Membrane 

translocation and activation of the Akt kinase in 

growth factor-stimulated hematopoietic cells. 

Leuk. Res. 21:1027-1031. 

Vanhaesebroeck, B. and Alessi, D.R. 2000. The 

PI3K-PDK1 connection: More than just a road 

to PKB. Biochem. J. 346:561-576. 

Yamada, K.M. and Araki, M. 2001. Tumor suppressor 

PTEN: Modulator of cell signaling, growth, 

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2382. 

Contributed by Roumen Pankov 

National Institute of Dental 

and Craniofacial Research 

National Institutes of Health 

Bethesda, Maryland 


Analyzing FAK and Pyk2 in Early 

Integrin Signaling Events 

Integrins are a family of heterodimeric α/β transmembrane receptors that bind to extracellular 

matrix proteins (ECM) proteins, and the signals generated by activated integrins 

regulate cell motility (Schwartz, 2001; Hynes, 2002). As integrins do not possess intrinsic 

signaling activity, integrin-stimulated signaling events promoting cell motility are 

initiated by integrin-associated kinases, e.g., focal adhesion kinase (FAK). FAK is one of 

several intracellular protein-tyrosine kinases (including the Src family, Pyk2, c-Abl, and 

Syk) that are activated by cell adhesion to ECM proteins such as fibronectin, collagen, or 

vitronectin. Integrin activation of FAK promotes increased FAK tyrosine phosphorylation 

and leads to the formation of an FAK-Src signaling complex (Schlaepfer et al., 1999; 

Schlaepfer and Mitra, 2004). Fibroblasts lacking FAK spread poorly and display migration 

defects in response to integrin stimuli (Ilic et al., 1995). Although the FAK-related 

kinase Pyk2 is expressed in FAK-null fibroblasts (FAK−/−), Pyk2 is not as effective 

in promoting FAK−/− cell motility as the FAK-Src signaling complex (Sieg et al., 

1998). This unit describes methods for culturing FAK+/+ and FAK−/− mouse embryo 

fibroblasts (MEFs; see Support Protocol 1) and stimulating these cells by replating onto 

fibronectin-coated dishes (see Basic Protocol 1). It also contains support protocols for 

cell starvation (see Support Protocol 2), preparation of protein cell lysates (see Support 

Protocol 3), and conditions and antibodies available for immunoprecipitation of FAK, 

Pyk2, and c-Src (see Support Protocol 4). 

Cell replating onto fibronectin activates FAK, and procedures are described for analyzing 

FAK-associated autophosphorylation activity (see Basic Protocol 2) with alternate protocols 

describing assays for measuring FAK-Pyk2 phosphorylation of a peptide substrate 

(see Alternate Protocol 1) and changes in c-Src-associated in vitro kinase activity (see 

Alternate Protocol 2). These catalytic measurement assays are complemented by methods 

using antibodies available for evaluating activity changes in FAK, Pyk2, or c-Src 

using immunoblotting techniques with phospho-specific antibodies (see Basic Protocol 

3). In addition, the biochemical analyses are complemented by cell biological methods to 

evaluate integrin-stimulated haptotaxis migration (see Basic Protocol 4) and time-lapse 

imaging to monitor wound closure motility in culture (see Basic Protocol 5). In these cell 

motility assays, transient plasmid expression vector transfection can be used to overexpress 

various proteins to test their effects on cell motility (Alternate Protocol 3). Finally, 

protocols are provided for visualizing activated FAK, Pyk2, and c-Src in cells plated onto 

extracellular matrix proteins (see Basic Protocol 6) with support methods for staining 

filamentous actin in cells (see Alternate Protocol 4), an easy means to visualize changes 

in cell shape and indirectly localize integrin-containing focal adhesions that are found at 

the ends of actin stress fibers. 

NOTE: All culture incubations should be performed in a humidified 37 ◦ C, 10% CO2 

incubator unless otherwise specified. Some media, e.g., DMEM, require higher levels of 


NOTE: All solutions and equipment coming into contact with cells must be sterile, and 

proper aeseptic technique should be used accordingly. 

Contributed by Joie A. Bernard-Trifilo, Ssang-Taek Lim, Shihe Hou, Dusko Ilic, and 

David D. Schlaepfer 


Copyright C○ 2006 by John Wiley & Sons, Inc. 

UNIT 14.7 





14.7.1 

Supplement 30

BASIC 

PROTOCOL 1 

Analyzing FAK 

and Pyk2 in Early 

Integrin Signaling 

14.7.2 

REPLATING ASSAYS FOR SIGNALING STUDIES 

Although anti-integrin antibodies can be used to facilitate integrin clustering, one of the 

strongest activators of integrin signaling is replating or passaging cells onto ECM-coated 

culture dishes. In the span of 1 hr, cells will rapidly bind to the ECM protein provided 

and will undergo cell spreading and morphology changes. Fibronectin binds to variety of 

α/β integrin pairs expressed on cells and is the strongest activator of FAK. Other ECM 

proteins can be used in replating assays, although the effective concentrations needed 

to stimulate integrin signaling are varied and dependent on cell type. This protocol describes 

the optimal conditions for replating FAK+/+ and FAK−/− cells on fibronectin to 

achieve maximal FAK or Pyk2-Src activation, respectively. It also provides a method for 

collecting protein lysates from these cells for further signaling studies. For comparative 

purposes, cell lysates are collected from adherent serum-starved cells, cells that have 

been held in suspension, cells replated onto fibronectin, and cells that have been replated 

onto a positively-charged ligand such as poly-L-lysine where rapid cell adhesion does 

not depend on integrin engagement. 

Materials 

Fibronectin (from bovine plasma; Sigma-Aldrich) 

Poly-L-lysine (mol. wt. 70,000 to 150,000 or 150,000 to 300,000; Sigma-Aldrich) 

Replating and migration medium (see recipe) 

FAK+/+ and FAK−/− MEFs, serum-starved (Support Protocol 2) 

Phosphate buffered saline (PBS; APPENDIX 2A) 

Trypsin/EDTA: 0.25% (w/v) trypsin/1 mM EDTA (Invitrogen) 

Trypsin inhibitor solution (see recipe) 

10-cm plastic tissue culture plates (Falcon) 

4 ◦ C incubator 

15-ml centrifuge tubes (Corning) 

Tabletop centrifuge 

10-ml transfer pipets, sterile 

50-ml conical centrifuge tubes (Corning) 

Light microscope 

Coat plates and prepare cells 

1a. To prepare fibronectin plates: Dilute fibronectin to 10 µg/ml in PBS and distribute 

5 ml per 10-cm plastic tissue culture plate. Incubate at 4 ◦ C overnight. 

1b. To prepare poly-L-lysine plates: Dilute poly-L-lysine to 50 µg/ml in PBS and distribute 

5 ml per 10-cm plastic tissue culture plate. Incubate at 4 ◦ C overnight. 

2. Prior to preparing cells, aspirate fibronectin or poly-L-lysine solutions, add 3 ml of 

replating and migration medium, aspirate, and place dishes at 37 ◦ C. 

The number of cell plates prepared depends on the number of experimental points to be 

evaluated. For example, to perform a time course of FAK activation, it is recommended 

that fibronectin and poly-L-lysine replating time points of 20, 40, 60, and 180 min be 

performed (see Support Protocol 3). 

3. Reserve one plate of serum-starved cells for a control. Wash serum-starved cells with 

PBS by adding 10 ml PBS and aspirating it off with a transfer pipet. Add 2.5 ml 

trypsin/EDTA per dish. Tap side of plates to facilitate cell release. 

4. Add 5 ml trypsin inhibitor solution warmed to 37 ◦ C. Transfer cells to a 15-ml 

centrifuge tube. Centrifuge 3 min at 750 × g, 21 ◦ C, in a tabletop centrifuge and 

aspirate supernatant. 


Replate cells 

5. Resuspend cells in 37 ◦ C replating and migration medium to 2 × 10 5 cells/ml. Keep 

cells in suspension at least 30 min and rotate tube gently at least every 5 min to keep 

cells from clumping, settling, and sticking to the plastic. 

It is convenient to pool all suspended cells into 50-ml conical centrifuge tubes such 

that subsequent cell distribution in replating assays is consistent. FAK−/− cells will 

sometimes clump together in suspension and forced pipetting to break apart clumps is 

recommended prior to replating. Keep additional cells in suspension during replating. 

6. Add 5 ml cell suspension to a 10-cm plate pre-coated with fibronectin or poly-L-lysine 

(from step 3) and incubate at 37 ◦ C. 

7. Periodically visualize replated cells under a microscope for changes in cell attachment 

and spreading. 

Maximal FAK activation occurs when cells have bound to fibronectin and are undergoing 

rapid cell spreading (between 20 and 60 min). Cells on poly-L-lysine will adhere, but will 

be very slow to spread. 

8. Proceed with Support Protocol 3 to prepare cell lysates of control cells (adherent 

serum-starved cells), experimental cells (replated on fibronectin or poly-L-lysine), 

and unplated serum-starved cells in suspension (45 min post replating). 

The unplated serum-starved cells in suspension and serum-starved experimental cells will 

be very important in correctly evaluating changes in FAK activation in immunoblotting 

(Basic Protocol 3) and in vitro kinase assays (Basic Protocol 2). 

GROWTH OF FAK+/+ AND FAK−/− FIBROBLASTS 

FAK-null MEFs were derived from an E8.0-day-old mouse embryo that had null mutations 

in genes for FAK and p53 (Ilic et al., 1995). Therefore, the cells are genetically 

FAK−/− p53−/−. Control FAK+/+ cells were derived in 1995 from an E8.0-day-old 

mouse embryo that had null mutations in p53 but not in FAK. Therefore, the cells are 

genetically FAK+/+ p53−/−. Both cell lines were derived from embryos of littermates 

obtained by crossing FAK+/− p53−/− female and FAK+/− p53−/− male mice. 

FAK+/+ and FAK−/− MEFs are grown on gelatin precoated plates because this was 

used during cell outgrowth from embryos. Although the fibroblasts are p53-null, the 

cells will become senescent (exhibited by an increase in cell size and the lack of cell 

proliferation) at high passage numbers. 

Materials 

Gelatin 

Phosphate buffered saline (PBS; APPENDIX 2A) 

FAK+/+ mouse embryo fibroblasts (ATCC CRL-2645) FAK−/− mouse embryo 

fibroblasts (ATCC CRL-2644): grown on plates as per ATCC product sheet 

Trypsin/EDTA: 0.25% (w/v) trypsin/1 mM EDTA (Invitrogen), 37 ◦ C 

Cell growth medium (see recipe), warmed to 37 ◦ C 

42 ◦ C water bath 

0.22-µm GP Express Plus filter membrane (Millipore) 

10-cm plastic tissue culture plates (Falcon) 

10-ml transfer pipets, sterile 

Light microscope 

15-ml conical centrifuge tubes (Corning) 

SUPPORT 

PROTOCOL 1 





14.7.3 


SUPPORT 

PROTOCOL 2 

Analyzing FAK 



14.7.4 

Prepare gelatin-coated plates 

1. Dissolve 0.5 g gelatin in 500 ml PBS to make 0.1% (w/v) solution. 

2. Heat in 42 ◦ C water bath to dissolve gelatin. 

3. Sterilize by passing through a Millipore 0.22-µm GP Express Plus filter membrane. 

Store up to 2 weeks at 4 ◦ C. 

4. Coat 10-cm plastic tissue culture plates with 5 ml sterile 0.1% gelatin solution and 

incubate 1 hr at 37 ◦ C 

5. Aspirate gelatin solution. Do not wash plates. 

Trypsinize cells 

6. Aspirate growth medium from a 10-cm plate of FAK−/− or FAK+/+ mouse embryo 

fibroblasts. 

7. Wash cells with 5 ml PBS. 

8. Add 2.5 ml warm trypsin/EDTA per plate. Tap side of plates to facilitate cell release. 

9. Incubate 2 to 3 min. Verify cell detachment under a microscope. 

FAK−/− cells adhere to the culture plates more firmly than FAK+/+ cells and will take 

longer to be released by trypsin/EDTA treatment. Senescent FAK−/− cells exhibit very 

strong adhesion to gelatin-coated dishes. 

10. Inactivate trypsin/EDTA by adding 5 ml cell growth medium. 

Replate cells 

11. Wash cells off plates by pipetting, and transfer them into 15-ml conical centrifuge 

tubes. 

12. Centrifuge cells in a tabletop centrifuge 3 min at 750 × g, 21 ◦ C. 

13. Aspirate supernatant and discard, retaining the pellet. 

14. Resuspend cells in fresh growth medium, count (see UNIT 1.1), and add growth medium 

to obtain the desired density. Add to gelatin-coated plates containing cell growth 

medium. 

Total volume is 8 to 10 ml medium per 10-cm plate. 

Consistent subcultivation is very important as cells that remain confluent for more than 

24 hr exhibit alterations in cell morphology and motility properties. In routine passage, 

FAK−/− cells may proliferate faster than FAK+/+ cells, so cell counting and normalization 

of cell numbers is required for equivalent experimental analyses. 

Both FAK−/− and FAK+/+ cells contain neomycin resistance genes and G418 

(neomycin) selection can be applied (500 µg/ml), although this is not absolutely required. 

15. Passage the cells when they are confluent. 

Split cells 1:10 to 1:15 every 3 days. Do not allow cells to become overly confluent, as 

this can result in a premature senescent phenotype. 

SERUM STARVATION OF CELLS 

For the stimulation and analysis of FAK activity and phosphorylation (Basic Protocols 1, 

2, and 3) and assays that involve cell replating onto extracellular matrix (ECM) proteins 

(Basic Protocols 4, 5, and 6), it is important to start with cells that are adherent and in 

a quiescent or nonproliferative state. This is accomplished by lowering the percent of 

serum to 0.5% in the growth medium. 


Additional Materials (also see Support Protocol 1) 

Starvation medium: prepared by making cell growth medium (see recipe) with FBS 

reduced to 0.5%. 

1. Passage cells (see Support Protocol 1) by plating 1 × 10 6 cells onto gelatin-coated 

10-cm dishes in growth medium. 

This is a subconfluent density, and it is important that cells do not reach confluency prior 

to initiation of cell motility analyses. 

2. After 24 hr, wash cells with PBS. 

3. Add 8 ml of starvation medium and incubate 16 to 24 hr at 37 ◦ C. Use cells immediately 

after starvation. 

Greater than 95% of FAK−/− and FAK+/+ cells should remain adherent under serumstarvation 

conditions. 

PREPARATION OF CELL LYSATES 

Activated FAK is associated with the cytoskeleton-rich fraction of cells that is not easily 

solubilized with low-detergent-containing buffers. The following lysate preclearing steps 

are used as the first step for immunoprecipitation, blotting, and in vitro kinase assays. 

All procedures are to be performed at 4◦C. Materials 

Experimentally treated, control adherent, and control suspended cells (Basic 

Protocol 1) 

Phosphate-buffered saline (PBS; APPENDIX 2A), 4 ◦ C 

RIPA cell lysis buffer (see recipe), 4 ◦ C 

50% (w/v) Sephadex G-100 (Sigma G100-120) slurry in PBS (APPENDIX 2A) 

Cell scraper (Corning) 

Refrigerated centrifuge 

1.5-ml microcentrifuge tubes 

1-ml disposable syringes 

21-G Luer-Lok needles 

Tube rotation device (e.g., Labquake, Barnstadt-Thermolyne) 

Microcentrifuge 

−80 ◦ C freezer (optional) 

1. Wash experimentally treated and adherent control cells with 10 ml cold PBS. 

2. At 45 min after replating the experimental cells (see Basic Protocol 1), centrifuge 

control suspended cells 3 min at 750 × g, 21 ◦ C. Resuspend cells in PBS and repeat 

centrifugation. 

3. Add 750 µl cold RIPA cell lysis buffer to each 10-cm plate (or 1 × 10 6 suspended 

cells). 

4. Scrape adherent cells using cell scraper and transfer to a 1.5-ml microcentrifuge 

tube. 

5. Using a 1-ml disposable syringe and 21-G Luer-Lok needle, pull cell lysate into 

syringe and shear cellular DNA by extrusion. Repeat at least five times. 

Be careful using the sharp needle and do not use too much force as this will result in 

excessive bubbling of the lysate. 

SUPPORT 

PROTOCOL 3 





14.7.5 


SUPPORT 

PROTOCOL 4 

Analyzing FAK 



14.7.6 

6. Add 100 µl 50% Sephadex G-100 slurry in PBS to each tube. Rotate tubes 10 min 

at 4 ◦ C. 

Sephadex G-100 beads will reduce DNA contamination and non-specific protein binding 

for subsequent immunoprecipitation analyses. 

7. Centrifuge 10 min at 16,000 × g, 4 ◦ C. 

8. Transfer supernatant (cleared total cell lysate) into new 1.5-ml microcentrifuge tubes. 

If the bead pellet is distributed along the sidewall of the microcentrifuge tubes, rotate 

tube direction 180◦ in centrifuge rotor and centrifuge an additional 5 min. 

9. Store cleared lysate indefinitely at −80 ◦ C or proceed with subsequent analyses. 

IMMUNOPRECIPITATION OF FAK, Pyk2, AND c-Src 

FAK is expressed at high levels in FAK+/+ fibroblasts. The FAK-related kinase, Pyk2, 

is expressed at low levels in FAK+/+ fibroblasts and at high levels in FAK−/− fibroblasts. 

Upon replating on fibronectin, both FAK and Pyk2 will form transient signaling 

complexes with the c-Src protein-tyrosine kinase. Replating of cells on fibronectin can 

also lead to the activation of c-Src through dephosphorylation of Tyr-527 located in 

the C-terminal domain of the regulatory region. This protocol describes standard immunoprecipitation 

methods used to isolate FAK, Pyk2, or multi-protein complexes with 

c-Src. 

Materials 

Primary antibody (see Table 14.7.1 and Table 14.7.2) 

Lysate of treated or control cells (Support Protocol 3) 

50% (w/v) protein A-agarose beads (fast flow immobilized protein A; Repligen, 

http://www.repligen.com) inPBS(APPENDIX 2A) 

50% (w/v) protein G-Plus-agarose beads (Calbiochem) in PBS (APPENDIX 2A) 

Triton lysis buffer (see recipe), 4 ◦ C 

HNTG buffer (see recipe), 4 ◦ C 


Tube rotation device (e.g., Labquake, Barnstadt-Thermolyne) 

Centrifuge 

Transfer pipets 

1. Add1to2µg of primary antibody to 0.75 ml lysate of treated or control cells in a 

microcentrifuge tube. 

If using cells from a frozen lysate, thaw at 37◦C then maintain at 4◦C. Use a FAK or Pyk2 antibody for subsequent analysis of FAK/Pyk2 kinase activity or 

phosphorylation. (Basic Protocol 2 or Alternate Protocol 1). 

Use an Src antibody for subsequent Src-associated analysis (Alternate Protocol 2). 

2. Rotate from 3 to 14 hr at 4 ◦ C. 

3. Add 30 µl of protein A– or protein G–bead slurry and rotate 60 min at 4 ◦ C. 

Protein A beads are used to capture rabbit polyclonal antibodies and protein G beads 

are used to bind goat polyclonal and mouse monoclonal antibodies (Table 14.7.1 and 

Table 7.2.1). 

4. Centrifuge beads 1 min at 1000 × g, 4 ◦ C. Remove supernatant with a transfer pipet. 

5. Add 800 µl Triton lysis buffer to beads, invert tube to suspend the bead pellet, and 

centrifuge 1 min at 1000 × g, 4 ◦ C. Remove supernatant with a transfer pipet and 

repeat step two more times. 


6. Add 800 µl of HNTG buffer, invert the tube to suspend the bead pellet, and centrifuge 

1 min at 1000 × g, 4 ◦ C. Remove supernatant with a transfer pipet and repeat step 

two more times. 

From this point, the immunoprecipitated samples can be analyzed in either the in vitro 

kinase assay (Basic Protocol 2) or resolved by SDS-PAGE (UNIT 6.1) and processed by 

immunoblotting (UNIT 6.2 and Basic Protocol 3). 

IN VITRO KINASE ASSAY 

Integrin binding to fibronectin leads to rapid cell spreading, and FAK, Pyk2, and c-Src 

tyrosine kinases become activated. The process can be characterized through changes associated 

with in vitro kinase activity. Assays are designed to detect auto-phosphorylation 

or the intramolecular phosphorylation of an added substrate such as a synthetic poly 

(Glu:Tyr) peptide for FAK and Pyk2, or the phosphorylation of a recombinant fragment 

of the FAK C-terminal domain by c-Src. 

Materials 

Immunoprecipitated samples (Support Protocol 4, step 6) 

FAK-Pyk2 kinase buffer (see recipe), 4 ◦ C 

Src kinase buffer (see recipe), 4 ◦ C 

10 µCi/µl [γ 32 P]ATP (>3,000 Ci/mmol; PerkinElmer) 

Magnesium/ATP cocktail (Upstate Biotechnology or see recipe) 

2× Laemmli SDS buffer (see recipe) 

Coomassie blue stain (see recipe) 

Molecular weight marker (Precision Plus, Bio-Rad) 

Destaining solution (see recipe) 

Microcentrifuge 


32 ◦ C water bath 

Plexiglas shielding 

Gloves 

Geiger counter 

Plexiglas box 

Micro tube cap locks (RPI 145063; http://www.rpicorp.com) 

Whatman 3MM filter paper 

Gel dryer 

Immobilon PVDF membrane (Millipore IPFL 000-10) 

Additional reagents and equipment for SDS-PAGE (UNIT 6.1), autoradiography 

(UNIT 6.3), and transfer of proteins to membranes for immunoblotting (UNIT 6.2) 



dispose of wastes in appropriately designated areas, following guidelines provided by 

the local radiation safety officer (also see APPENDIX 1A). 

1. Wash the immunoprecipitated samples (Support Protocol 4, step 6) with 

800 µl cold FAK-Pyk2 kinase buffer two times, centrifuging 2 min at 16,000 × 

g, 4 ◦ C, each time. 

2. Aspirate supernatant, but leave a small amount above pellet (∼20 to 30 µl). 

3. For FAK-Pyk2 autophosphorylation assays, add 2.5 µlof10µCi/µl[γ 32 P] ATP and 

5 µl magnesium/ATP cocktail to the immune complexes. Slightly flick tubes to mix 

and place in 32 ◦ C water bath for 15 min. 

BASIC 

PROTOCOL 2 





14.7.7 


Analyzing FAK 



14.7.8 

Figure 14.7.1 Measurements of Pyk2 of FAK-associated in vitro kinase activity. Lysates from 

serum-starved (attached), suspended (S) fibronectin-plated (FN), and poly-L-lysine-plated (PL) 

FAK−/− cells (A and B) or FAK+/+ cells (C and D) were prepared and divided into equal aliquots 

for either Pyk2 immunoprecipitates (IPs) or FAK IPs, respectively. (A) Pyk2 IPs were labeled by 

the addition of [γ- 32 P]ATPinaninvitrokinase(IVK)assay. 32 P-labeled proteins were transferred 

to membranes and visualized by autoradiography. (B) The same membrane shown in (A) was cut 

and analyzed by either anti-Pyk2 or anti-Src family PTK blotting. (C) FAK IPs were labeled by the 

addition of [γ- 32 P]ATP in an IVK assay and the same membrane (D) was cut and analyzed by 

either anti-FAK or anti-Src family protein-tyrosine kinase (PTK) immunoblotting. Molecular weight 

standards are indicated in kDa to the left. (previously published in the EMBO Journal; Siegetal., 

1998) 


CAUTION: Place Plexiglas shielding around gel area to prevent radiation exposure. 

Remember to check gloves and work areas with a Geiger counter after the kinase assay 

is done. Use a Plexiglas box for moving radioactive materials within the laboratory and 

dispose of radioactive waste into designated storage according to institutional requirements. 

4. Add 50 µl 2× Laemmli SDS buffer to stop kinase reactions. Secure 1.5-ml tubes 

with micro tube cap locks. 

5. Boil samples for 3 min at 100 ◦ C. 

6. Microcentrifuge agarose beads 2 min at 16,000 × g, 21 ◦ C. Load supernatants and 

molecular weight marker onto a 7.5% SDS-PAGE gel (see UNIT 6.1). 

For autoradiography 

7a. After gel is run (∼4 hr), cut one top corner to mark orientation. Place gel in Coomassie 

blue stain for 1 hr and then destain gel for 6 to 12 hr in multiple changes of destaining 

solution on a shaking platform. Wash a final time in water. 

8a. Place gel onto Whatman 3MM filter paper and dry using a gel dryer. 

9a. Expose gel to film or phosphor-imager screen (UNIT 6.3) to quantify 32 P incorporated 

into FAK (∼116 kDa), Pyk2 (∼110 kDa) or associated c-Src (∼60 kDa). 

See Fig. 14.7.1 A and C. 

For autoradiography and subsequent immunoblotting 

7b. Transfer the gel to a PVDF membrane via semidry electrophoretic transfer (see 

UNIT 6.2). 

8b. Stain min with Coomassie blue stain. Destain 10 min in destaining solution and wash 

a final time in water. 

9b. Place membrane between protective plastic sheets and expose gel to film or phosphorimager 

screen to quantify 32 P incorporated into FAK (∼116 kDa), Pyk2 (∼110 kDa) 

or associated c-Src (∼60 kDa). See UNIT 6.3 for more information. 

In this manner, subsequent immunoblotting (see Basic Protocol 3) can be performed on 

the kinase reactions to verify the amount of FAK or Pyk2 in the immune complex or the 

amount of c-Src associated with the activated FAK and/or Pyk2 complex reaction (see 

Fig. 14.7.1B and D). 

FAK-PYK2 POLY GLU:TYR PHOSPHORYLATION 

This procedure is used to measure the autophosphorylation of FAK/Pyk2 (as in Basic 

Protocol 2) combined with the transphosphorylation of an added generic substrate 

peptide. 

Additional Materials (also see Basic Protocol 2) 

Immunoprecipitated samples (see Support Protocol 4, step 6) 

10 mg/ml poly(Glu:Tyr; 4:1) mol. wt. 20,000 to 50,000: sodium salt 

(Sigma-Aldrich) prepared in PBS and stored up to 2 years at −20 ◦ Cin1-ml 

aliquots 

Magnesium/ATP cocktail (Upstate Biotechnology or see recipe) 

10 µCi/µl [γ 32 P]ATP (>3,000 Ci/mmol; PerkinElmer) 

FAK-Pyk2 kinase buffer 

0.75% phosphoric acid 

100% acetone 

Scintillation fluid (Sigma) 

2 × 2–cm Whatman 3MM filter paper squares 

ALTERNATE 

PROTOCOL 1 





14.7.9 


ALTERNATE 

PROTOCOL 2 

Analyzing FAK 



14.7.10 

Conical 50-ml centrifuge tube 

Scintillation vials 

Scintillation counter 

Additional reagents and equipment for preparing immunoprecipitated samples 

(Support Protocol 4) 





1. Wash the immunoprecipitated samples (from Support Protocol 4, step 6) with 800 µl 

cold FAK-Pyk2 kinase buffer two times, centrifuging 2 min at 16,000 × g, 4 ◦ C, each 

time. 

2. Remove supernatant, but leave a small amount above pellet (∼20 to 30 µl). 

3. Add 5 µl 10 mg/ml poly(Glu:Tyr), 5 µl magnesium/ATP cocktail, and 2.5 µl of 

10 µCi/µl [γ 32 P]ATP (25 µCi) to the immunoprecipitated samples. Slightly flick 

tubes to mix and place in 32 ◦ C water bath for 15 min. Include a negative control for the 

phosphorylation assay, substituting 35 µl kinase buffer for the immunoprecipitated 

samples. 





The negative control will measure the level of background binding of [ 32P ATP] to the 

assay squares. 

The total volume of immune complex reaction is ∼50 µl yielding a final concentration of 

50 µM ATP from the magnesium/ATP cocktail and 1 mg/ml poly Glu:Tyr. 

4. Transfer a 10-µl aliquot onto the center of a labeled (with a #2 pencil) 2 × 2–cm 

Whatman filter paper square. 

There should be enough volume for triplicates for each immune complex reaction. 

5. Allow the radiolabeled substrate to bind to filter paper for at least 1 min. Transfer 

filter papers to a 50-ml conical tube containing 40 ml 0.75% phosphoric acid. Gently 

invert tube to wash the assay squares for 5 min. Repeat three times and discard liquid 

as radioactive waste. 

Ten or more assays squares can be washed per 50-ml tube. 

6. Wash assay squares once with 40 ml of 100% acetone for 5 min. Discard liquid waste 

and transfer assay squares to scintillation vial. Add scintillation fluid and evaluate 

using a scintillation counter. 

For information on creating a standard curve, see http://www. perkinelmer.com. 

MEASUREMENTS OF Src-ASSOCIATED KINASE ACTIVITY 

As c-Src contains both stimulatory (Tyr-416) and inhibitory (Tyr-527) phosphorylation 

sites that act to regulate catalytic activity, measurements to analyze changes in 

c-Src-associated kinase activity are performed by c-Src immune complex phosphorylation 

of an FAK C-terminal domain fragment that contains the known c-Src phosphorylation 

sites, FAK Tyr-861 and FAK Tyr-925. 



10 mg/ml GST-FAK 853-1052 (Schlaepfer lab) 

Enolase (Sigma-Aldrich E-0379), optional 

50 mM HCl, optional 

1 M PIPES, pH 7, optional 

30◦C water bath, optional 





1. Wash the immunoprecipitated samples (from step 6 in Support Protocol 4) with 

800 µl cold FAK-Pyk2 kinase buffer two times, centrifuging 2 min at 16,000 × g, 

4 ◦ C, each time. 

2. Remove supernatant, but leave a small amount above pellet (∼20 to 30 µl) 

3. Add 5 µl of 10 mg/ml GST-FAK 853-1052, 5 µl magnesium/ATP cocktail, and 2.5 µl 

of 10 µCi/µl[γ 32 P]ATP (25 µCi) to the immunoprecipitated samples. Slightly flick 

tubes to mix and place in 32 ◦ C water bath for 15 min. 





The total volume of immune complex reaction is ∼50 µl yielding a final concentration of 

50 µM ATP from the magnesium/ATP cocktail and 1 mg/ml GST-FAK 853-1052. 

Alternatively, acid-denatured enolase may be used as a substrate. Enolase must be 

freshly prepared: Add 10 µl of enolase (1 mg/ml) to 2 µl of 50 mM HCl and incubate 

10 min at 30◦C. Neutralize with 2 µl 1 M PIPES, pH 7.0, and use 2 µl acid-denatured 

enolase per kinase reaction. 

4. Proceed with Basic Protocol 2, steps 4 through 9 to perform SDS-PAGE and autoradiography 

or autoradiography and immunoblotting. 

IMMUNOBLOTTING WITH FAK/PYK2 PHOSPHO-SPECIFIC ANTIBODIES 

While in vitro kinase assays will reflect changes in catalytic activity, immunoblotting 

with FAK/PYK2 phospho-specific antibodies can be used to assess specific phosphorylation 

site modification(s) within the kinases. For example, FAK and Pyk2 first become 

phosphorylated at their autophosphorylation sites Y397 and Y402 respectively. Upon 

phosphorylation and binding of c-Src in an active signaling complex, c-Src can phosphorylate 

other sites on FAK and Pyk2. Additionally, there are several serine-threonine sites 

on FAK that can be phosphorylated by other cellular kinases. Finally, phospho-specific 

antibodies have been developed to indirectly measure the activation state of c-Src. 

Materials 

2× Laemmli SDS buffer (see recipe) 

Immunoprecipitated samples (Support Protocol 4, step 6) 

100% and 20% methanol 

Transfer buffer (see UNIT 6.2) 

EZBlue gel staining reagent (Sigma) 

BSA blocking buffer (see recipe) 

Primary antibody (see Tables 14.7.1 and 14.7.2) 

BASIC 

PROTOCOL 3 





14.7.11 


Analyzing FAK 



14.7.12 

Tris-buffered saline with Tween (TBST, see recipe) 

Secondary antibody: Horseradish peroxidase (HRP)-conjugated appropriate 

species (Pierce) 

Enhanced chemiluminescence (ECL) western detection solution (Amersham RPN 

2132) 

Western blot stripping buffer (see recipe) 

Immobilon PVDF membrane (Millipore) 

Rotating shaker 

Additional reagents and equipment for gel electrophoresis (UNIT 6.1) and 

electrophoretic transfer (UNIT 6.2) 

Perform gel electrophoresis 

1. Add 50 µl of2× Laemmli SDS buffer to the immunoprecipitated samples from 

Support Protocol 4, step 6 and boil samples 3 min at 100 ◦ C. 

2. Centrifuge 2 min at 16,000 × g, 21 ◦ C. Using the supernatants perform SDS-PAGE 

(UNIT 6.1), including molecular size markers. 

Transfer proteins to PVDF membrane 

3. Transfer proteins to a PVDF membrane using an electrophoretic transfer unit 

(UNIT 6.2). 

PVDF membrane should be prepared by soaking in 100% methanol followed by the 

appropriate transfer buffer before use. 

4. After transfer is complete, stain protein bands with EZBlue gel staining reagent. 

Rinse membrane in water and destain in 20% methanol until background staining 

disappears. 

5. Photocopy membrane for record keeping purposes and to note positions of protein 

size markers. 

Probe membrane 

6. Place membrane in BSA blocking buffer for 1 hr at room temperature. 

7. Remove blocking buffer and incubate membranes with primary antibodies 3 hr at 

room temperature or overnight at 4 ◦ C. 

A 1:1000 dilution (∼0.5 µg/ml in BSA blocking buffer) is a good starting point for using 

an antibody for the first time. 

8. Wash membranes by rotational shaking with TBST for 10 min. Repeat three times 

using fresh TBST. 

9. Dilute secondary antibody 1:5000 to 1:10,000 in TBST (e.g., 5 µl in 15 ml) and 

incubate membranes with the antibody 1 hr at room temperature. 

HRP-conjugated secondary antibodies commonly used are: goat anti-mouse IgG, Protein 

A, donkey anti-rabbit IgG, and mouse anti-goat IgG. 

10. Wash membranes by rotational shaking with TBST for 10 min. Repeat three times 

using fresh TBST. 

11. Incubate in 5 ml ECL solution for 5 min at 21 ◦ C. Expose to film 30 sec to 10 min to 

achieve optimal exposure. 

Although many phospho-specific antibodies can be used to probe FAK, Pyk2, or 

c-Src activation states in whole cell lysates, multiple immunoreactive bands detected 

can complicate the interpretation of results. This is not a problem when analyzing FAK, 

Pyk2, or c-Src by immunoprecipitation. 


Table 14.7.1 Commercially Available Antibodies to FAK, Pyk2, and c-Src a 

Antibody Company Source Application Catalog number 

FAK, clone 4.47 Upstate mouse (mAb) WB, IP, IC, IH 05-537 

FAK clone 2A7 Upstate mouse (mAb) IP, IC 05-182 

FAK Upstate rabbit WB, IP, IH 06-543 

FAK, BC3 Upstate rabbit IP, IC 06-446 

FAK BioSource rabbit IP, WB AHO0502 

FAK BD PharMingen rabbit WB, IP 556368 

FAK BD Transduction mouse (mAb) WB, IP, IC, IH 610087 

FAK Cell Signaling rabbit WB, IH 3285 

FAK BioSource rabbit WB, IP, IC AH0502 

FAK BioSource rabbit WB, IP, IC AMO0672 

FAK Chemicon rabbit IP AB1605 

FAK Chemicon mouse (mAb) WB, IP, IC MAB2156 

FAK (H-1) Santa Cruz mouse (mAb) WB, IP, IC, IH sc-1688 

FAK (A-17) Santa Cruz rabbit WB, IP, IC sc-557 

FAK (C-20) Santa Cruz rabbit WB, IP, IC sc-558 

FAK (C-903) Santa Cruz rabbit WB, IP, IC, IH sc-932 

Pyk2 Upstate rabbit WB, IP, IC 06-559 

Pyk2, clone 74 Upstate mouse (mAb) WB, IP 05-488 

Pyk2 Upstate rabbit WB, IP, IC 07-437 

Pyk2 Cell Signaling rabbit WB, IP 3292 

Pyk2 BD Transduction mouse (mAb) WB, IP, IC, IH 610548 

Pyk2 (H-102) Santa Cruz mouse (mAb) WB, IP, IC sc-9019 

Pyk2 (N-19) Santa Cruz rabbit WB, IP, IC sc-1514 

Pyk2 (C-19) Santa Cruz rabbit WB, IP, IC sc-1515 

c-Src, clone GD11 Upstate mouse WB, IP 05-184 

c-Src, clone N6L Upstate rabbit (mAb) WB 05-889 

c-Src, clone NL19 Upstate rabbit (mAb) WB, IP 05-772 

c-Src BioSource mouse (mAb) WB AHO1152 

c-Src BioSource rabbit WB 44-655G 

c-Src BioSource rabbit WB 44-656G 

c-Src Chemicon sheep WB, IP CB769 

c-Src, clone 36D10 Cell Signaling rabbit (mAb) WB, IP, IC, IH 2109 

c-Src, clone L4A1 Cell Signaling mouse (mAb) WB, IP 2110 

c-Src Cell Signaling rabbit WB, IP, IC, IH 2108 

c-Src (H-12) Santa Cruz mouse (mAb) WB, IP, IC sc-5266 

c-Src (B-12) Santa Cruz mouse (mAb) WB, IP, IC sc-8056 

c-Src (N-16) Santa Cruz rabbit WB, IP, IC sc-19 

c-Src (Src 2) Santa Cruz rabbit WB, IP, IC sc-18 

aAbbreviation: mAb, monoclonal antibody; WB, western (immuno)blot; IP, immunoprecipitation; IC, immunocytochemistry; 

IH, immunohistochemistry. 





14.7.13 


Analyzing FAK 



14.7.14 

Table 14.7.2 Commercially Available Phospho-Specific Antibodies to FAK, Pyk2, and c-Src a 


FAK pY397 BioSource rabbit WB, IC, IH 44-624G 

FAK pY397 

clone 141-9 

BioSource rabbit (mAb) WB, IC 44-625G 

FAK pY407 BioSource rabbit WB, IC, IH 44-650G 

FAK pY576 BioSource rabbit WB 44-652G 

FAK pY577 BioSource rabbit WB, IC 44-614G 

FAK pS722 BioSource rabbit WB 44-588 

FAK pS732 BioSource rabbit WB 44-590G 


FAK pY861 BioSource rabbit WB, IC 44-626G 


FAK pY397 

clone 14 

BD/Transduction mouse (mAb) WB, IC 611722 

FAK pY397 

clone 18 

BD/Transduction mouse (mAb) WB, IC 611806 

FAK 

pY576/pY577 

Cell Signaling rabbit WB 3281 

FAK pY397 Santa Cruz rabbit WB, IC sc-11765-R 


FAK pY407 Santa Cruz goat WB, IC sc-16664 



FAK 

pY576/pY577 

Santa Cruz rabbit WB, IC sc-21831-R 

FAK 

pY576/pY577 

Santa Cruz goat WB, IC sc-21831 

FAK pS722 Santa Cruz goat WB, IC sc-16662 


FAK pS910 Santa Cruz goat WB, IC sc-16666 


FAK pY397 Chemicon mouse (mAb) WB, IC MAB1144 

FAK pY397 Upstate rabbit WB, IC 07-012 

FAK pY576 Upstate rabbit WB, IC 07-157 

Pyk2 pY402 BioSource rabbit WB, IH 44-618G 

Pyk2 pY579 BioSource rabbit WB, IC 44-632G 

Pyk2 

pY579/pY580 

BioSource rabbit WB 44-636G 

Pyk2 pY580 BioSource rabbit WB 44-634G 

Pyk2 pY881 BioSource rabbit WB, IH 44-620 

Pyk2 pY402 Cell Signaling rabbit WB, IP 3291 

continued 


Table 14.7.2 Commercially Available Phospho-Specific Antibodies to FAK, Pyk2, and c-Src a , 

continued 


>Pyk2 pY402 Santa Cruz rabbit WB, IC sc-11767-R 

Pyk2 pY579 Santa Cruz goat WB, IC sc-16822 

Pyk2 

pY579/pY580 

Santa Cruz goat WB, IC sc-16824 



Pyk2 pY402 

clone RR102 

Upstate mouse (mAb) IP 05-679 

Active Src 

clone 28 

BioSource mouse (mAb) WB, IC, IH AHO0051 

Src pY416 BioSource rabbit WB, IC 44-660G 

Src pY527 BioSource rabbit WB, IH 44-662G 

Src pY527 

clone 31 

BD/Transduction mouse (mAb) WB 612668 

Src pY416 Cell Signaling rabbit WB, IC, IH 2101 

Src pY416 

clone 7G9 

Cell Signaling mouse (mAb) WB, IP 2102 

Non-phospho 

Src pY527 

Cell Signaling rabbit WB 2107 

Src pY527 Cell Signaling rabbit WB, IH 2105 

Src pY527 Santa Cruz goat WB, IC sc-16846 

Src pY416 

clone 2N8 

Upstate rabbit (mAb) WB 05-857 

Src pY416 

clone 9A6 

Upstate mouse (mAb) WB 05-677 

aAbbreviations: mAb, monoclonal antibody; WB, western (immuno)blot; IP Immunoprecipitation; IC, immunocytochemistry; 

IH, immunohistochemistry. 

Reprobe membranes 

12. For sequential probing of membranes with different antibodies, strip bound antibodies 

from membranes by incubating 15 min in western blot stripping buffer at 65 ◦ C. 

Wash membrane thoroughly with water. 

13. Repeat immunoblotting procedure from step 4 to step 9. 

Blots can be stripped two to three times to analyze FAK and Pyk2 immune complexes 

with various phospho-specific antibodies. It is recommended that a different species of 

primary antibody be used (rabbit, mouse, or goat) in the subsequent blotting analyses 

to ensure that signals detected are not from the first set of antibodies used to probe the 

membrane. 

HAPTOTAXIS MOTILITY ASSAY: MATRIX-STIMULATED MIGRATION 

FAK is rapidly activated via tyrosine phosphorylation and binds directly to c-Src, resulting 

in the formation of a FAK-Src signaling complex that leads to the activation of various 

downstream signaling cascades. In the preceding protocols, emphasis was placed on 

biochemical evaluations of FAK, Pyk2, or Src activation or phosphorylation after cell 

binding to extracellular matrix proteins. In addition to biochemical signaling changes 

BASIC 

PROTOCOL 4 





14.7.15 


Analyzing FAK 



14.7.16 

inside cells, cell binding to matrix proteins such as fibronectin can result in enhanced cell 

spreading and the initiation of cell migration. Notably, FAK−/− cells spread poorly and 

exhibit refractory motility responses in response to a fibronectin stimulus. Although it has 

been shown that FAK reexpression within FAK−/− cells can reverse the motility defects, 

the molecular pathways through which FAK promotes cell motility and the inability of 

Pyk2 in FAK−/− cells to function as a replacement for FAK remain active areas of 

research. There are various assays whereby cell motility can be measured. To specifically 

evaluate the contribution of integrin signaling, haptotaxis (a directed response of cells 

in a gradient of adhesion) motility assays are performed in the absence of serum where 

FAK signaling plays important roles in promoting efficient cell movement towards the 

extracellular matrix. 

Materials 

FAK−/− and FAK+/+ cells, subconfluent (see Support Protocol 1) 

Human plasma fibronectin (Sigma-Aldrich F2006): 2 to 10 µg/ml in replating and 

migration medium (see recipe), 37 ◦ C 

BSA-coating control: 10 µg BSA/ml in replating and migration medium (see 

recipe) 

Trypsin/EDTA: 0.25% (w/v) trypsin/1 mM EDTA (Invitrogen) 

Trypsin inhibitor solution (see recipe) 

PBS ++ : phosphate-buffered saline (PBS; APPENDIX 2A) with 0.1 g/liter CaCl2 and 

0.5 mM MgCl2 

Cell fixative: PBS with 1.85% (v/v) formaldehyde and 0.05% (v/v) glutaraldehyde 

10% (w/v) crystal violet stain (Sigma) in ethanol 

0.1 M sodium borate, pH 9.0 

Parafilm 

Millicell PCF chamber inserts, 8-µm pore (Millipore PITP01250) 

Flat-tipped forceps 

15-ml centrifuge tube 

Tabletop centrifuge 

Hemacytometer 

24 well-tissue culture dishes (Costar) 

Cotton swabs 

Inverted light microscope 

Spectrophotometer with A600 capability, optional 

Additional reagents and equipment for serum-starving cells (Support Protocol 2) 

and counting cells (UNIT 1.1) 

Starve cells 

1. Twenty-four hr before the motility assay, serum starve subconfluent FAK−/− and 

FAK+/+ cells (see Support Protocol 2). 

Cells should be subconfluent during starvation. High-density or contact-inhibited cells 

do not respond well to motility stimuli during the short time period of the assay. 

Prepare assay chamber 

2. Pipet 100 µl of human plasma fibronectin onto Parafilm. Carefully place Millicell 

PCF chamber inserts one at a time on top the fibronectin-containing droplets. Prepare 

three chambers per experimental point and include a BSA-coating control. 

The use of flat tipped forceps is best for the handling of individual chambers. 

Be sure to cover the entire membrane and to avoid bubbles as this will result in poor 

ligand coverage. This method is designed to coat just the under surface of the Millicell 

chamber with fibronectin. 


3. Incubate chambers 2 hr at room temperature. Cover the Parafilm and chambers on 

the bench top with a plastic container to avoid evaporation. 

4. Rinse each chamber gently in PBS and let dry for at least 10 min at room temperature. 

Prepare cells 

5. Wash serum-starved cells with 10 ml PBS and aspirate off. Add 2.5 ml trypsin/EDTA 

per dish. Tap side of plates to facilitate cell release. 

6. Add 5 ml trypsin inhibitor solution warmed to 37 ◦ C. Transfer cells to 15-ml centrifuge 

tube. Centrifuge 3 min at 750 × g,21 ◦ C, in a tabletop centrifuge and aspirate 

supernatant. 

7. Resuspend cells to ∼2 × 10 5 cells/ml in 37 ◦ C replating and migration medium. Keep 

in suspension at least 30 min and rotate tube at least every 5 min to keep cells from 

clumping, settling, and sticking to the plastic. 

8. During incubation period, count cells using a hemacytometer (UNIT 1.1) and dilute 

a suspension of cells to a density of 33 × 10 4 cells/ml in replating and migration 

medium 

Perform migration assay 

9. Add 400 µl replating and migration medium to each experimental well of a 24-well 

tissue culture dish. Carefully place chambers into wells making sure there are no 

bubbles. 

10. Add 300 µl cell suspension to each chamber (total of 1 × 10 5 cells). 

11. Incubate 3 to 4 hr at 37◦C with 10% CO2. 

Fibroblasts will show maximal migration within this period. 

Variations of this assay can be used to examine different types of motility. For example, 

chambers can be coated on the top and bottom to study random motility. Additionally, 

growth factors can be added to the lower chamber to investigate cell chemotaxis responses. 

Fix, stain, and count cells 

12. Fill two 24-well tissue culture plates with 0.5 ml PBS ++ and one with 0.5 ml cell 

fixative. Transfer chambers to PBS ++ and incubate 2 min to wash. Move chambers 

to fixative for 5 min then into fresh PBS ++ for 2 min. 

13. Dilute 10% crystal violet stain stock 1:100 in 0.1 M sodium borate, pH 9.0. Place 

chambers into 0.5 ml of 0.1% crystal violet for 30 min. 

14. Remove chambers from stain and wash repeatedly in a large volume of water (e.g., 

in a 500 ml beaker). 

15. Use a cotton swab to wipe excess cells and stain from the top side of the chamber 

membrane while taking care not to touch the bottom side of the chamber membrane. 

Let chambers dry overnight. 

Chambers are now ready for analysis. Cells that migrate to the bottom of the chamber 

towards the ligand will be stained purple with crystal violet while nonmigrating cells on 

the upper surface have been removed. 

16. Count cells using a standard 10× objective on an inverted microscope. Use the lid 

of the 24-well plate to support the chamber. Count fields at 2, 4, 6, 8, 10, and 12 

o’clock positions and in the center of the membrane. 

If there are too many cells per field to be accurately counted, the crystal violet stain can be 

eluted from the membrane in 10% acetic acid and values can be obtained by measuring 

light absorption in a spectrophotometer at 600 nm (A600). 





14.7.17 


ALTERNATE 

PROTOCOL 3 

Analyzing FAK 



14.7.18 

Motility values are determined by the average number of cells per field when counting 

three chambers per experimental point. The BSA-coated chambers should contain less 

than 1% of the total as migratory cells. 

ANALYZING CELL MOTILITY USING PLASMID-TRANSFECTED CELLS 

The haptotaxis protocol is a good way to evaluate integrin-stimulated signaling events 

affecting cell migration. In many instances, there is a need to test the role of particular 

signaling proteins in altering cell motility responses. This is best accomplished through 

the transient transfection of cells with plasmid expression vectors combined with the 

cotransfection of a marker such as β-galactosidase (lac Z) to identify the plasmidtransfected 

cells, the assumption being that the cells that have been transfected with the 

lac Z plasmid and show β-galactosidase activity have also taken up the cotransfected 

FAK plasmid. This protocol is used to show that FAK re-expression can rescue the 

motility defects of FAK−/− cells. 


pcDNA3.1 FAK (contact Schlaepfer lab) 

FAK−/− cells (see Support Protocol 1) 

pcDNA3.1 LacZ (Invitrogen) 

Phosphate-buffered saline (PBS; APPENDIX 2A) 

PBS ++ : phosphate-buffered saline (APPENDIX 2A) with 0.1 g/liter CaCl2 and 0.5 mM 

MgCl2 

Opti-MEM I reduced-serum medium (Invitrogen) 

PLUS reagent (Invitrogen) 

Lipofectamine (Invitrogen) 

Cell growth medium (see recipe) containing 20% (v/v) FBS (instead of 10%) 

Starvation medium: Cell growth medium (see recipe) with FBS reduced to 0.5% 

Lac Z staining solution (see recipe) 

Additional reagents and equipment for bacterial transformation (Seidman et al., 

1997), plasmid miniprep (Engebrecht et al., 1991), and DNA quantification 

(APPENDIX 3D) 

Prepare plasmids 

1. Prepare high purity supercoiled plasmid DNA from bacteria (see Seidman et al., 

1997; Engebrecht et al., 1991; and APPENDIX 3D). Combine 5 µg of pCDNA3.1 FAK 

with 3 µg pcDNA3.1 LacZ for each 10-cm plate. 

Make sure the DNA is sterile by using alcohol precipitation. 

Maximum amount of DNA per 10-cm plate transfection is 8.0 µg. 

Prepare cells 

2. Plate 7.5 × 10 5 FAK−/− cells onto gelatin-coated 10-cm plates (see Support Protocol 

1) and let grow overnight at 37 ◦ C. 

3. Aspirate medium and wash once with PBS to remove serum. 

4. Add 4 ml of Opti-MEM I reduced-serum medium and place cells in the 37 ◦ C 

incubator for 30 to 45 min. 

Prepare transfection mix 

5. Aliquot 0.5 ml Opti-MEM I reduced-serum medium into two 1.5-ml microcentrifuge 

tubes (label tubes A and B). 

6. Add 8 µg plasmid DNA (FAK plus LacZ) to tube A and resuspend by tapping with 

your fingers. 


7. Add 48 µl PLUS reagent to the DNA in tube A and mix by tapping. 

8. Add 32 µl Lipofectamine reagent to tube B. Mix by tapping and incubate for 15 min. 

9. Add contents of tube B to tube A and mix by tapping. Incubate for 30 min at 21◦C. IMPORTANT NOTE: Do not mix by pipetting at this step. 

Transfect cells 

10. Add the DNA/lipid complexes from step 9 to the cells from step 4 and allow cell 

transfection to proceed for 5 hr at 37 ◦ C in the 10% CO2 incubator. 

11. Add 5 ml cell growth medium containing 20% FBS to stop transfection. Continue 

incubation to 24 hr without changing the medium. 

Prepare cells for motility assays 

12. To set up cells for motility assays, wash with 10 ml PBS and place in 10 ml serum 

starvation medium and incubate overnight at 37 ◦ C in 10% CO2. 

13. Use cells for motility experiments 48 hr after transfection to ensure good protein 

expression from plasmids. 

14. Proceed with the motility assay (Basic Protocol 4, steps 2 to 11). 

15. Save excess cells not used in the motility assay to make protein lysates (Support 

Protocol 3) and to analyze by SDS-PAGE (UNIT 6.1) followed by immunoblotting 

(Basic Protocol 3 and UNIT 6.2). 

This is required in order to verify transient protein overexpression (i.e., FAK and Lac Z) 

in the population of cells used for the motility assay. 

Fix and count cells 

16. After haptotaxis assays have proceeded for 3 to 4 hr, fill two 24-well tissue culture 

plates with 0.5 ml PBS ++ and one with 0.5 ml cell fixative. Transfer chambers to 

PBS ++ and incubate for 2 min (wash). Move chambers to fixative for 5 min then 

into fresh PBS ++ for 2 min. Repeat final PBS wash. 

It is important to not leave cells in fixative for too long as this will result in less 

β-galactosidase activity. 

17. Use a cotton swab to wipe excess cells from the top side of the chamber membrane 

while taking care not to touch the bottom side of the chamber membrane. Do not let 

chambers get too dry. 

18. Add 0.5 ml Lac Z staining solution to wells of a 24-well tissue culture plate and 

incubate chambers 2 to 12 hr at 37 ◦ C or until blue-green color is observed. 

19. Rinse chambers by gently immersing them in PBS and enumerate the blue β-gal 

positive cells under light microscopy. 

20. To store plates fix chambers 10 min with 1 ml 10% formalin in PBS at room 

temperature, rinse with PBS, and store in PBS up to 1 week at 4 ◦ C. 

It is assumed that the cells that have been transfected with the Lac Z plasmid and show 

β-galactosidase activity have also taken up the cotransfected FAK plasmid. If the protein 

of interest is tagged with a fluorescent marker such as green fluorescent protein, then the 

direct identification of transfected cells is facilitated, making it possible to observe the 

behavior of such cells in time-lapse imaging studies (see Basic Protocol 5). 





14.7.19 


BASIC 

PROTOCOL 5 

Analyzing FAK 



14.7.20 

SCRATCH-WOUND HEALING ASSAY WITH TIME-LAPSE IMAGING 

Wound healing assays (UNIT 12.4) are ideal for visualizing cellular dynamics during stimulated 

migration. In particular, time-lapse imaging allows for analysis of lamellipodia 

extension and membrane ruffling into the wounded area. Wound healing assays do not, 

however, address directional motility because the cells are plated at high density and 

movement is limited into the wound. Random migration assays can be preformed to 

address directional motility by plating cells at low density and following the imaging 

protocol outlined below. 

Materials 

Extracellular matrix molecule (ECM) of interest (e.g., 2 µg fibronectin/ml PBS) 

70% confluent 24-hr serum-starved cells (see Support Protocol 2) 

DMEM with and without 0.5 µg/ml mitomycin-C 


Serum, optional 

Mitomycin-C (Sigma) 

Medium 199 (Invitrogen) with 0.5 µg/ml mitomycin-C 

Mineral oil (Sigma) 

25-mm glass coverslips (1 oz., Fisher) and 6-well tissue culture plates or 

35-mm Bioptechs delta-T dishes (Fisher) 

Forceps 

1- to 10-µl micropipet tips 

Transfer pipets, sterile 

Inverted microscope with 20× objective, 37 ◦ C heated stage, and 

acquisition/analysis software (e.g., Improvision Openlab; 

https://www.improvision.com) 

Etched-grid coverslips (Bellco), optional 

Coat coverslips 

1. Place 25-mm round glass coverslips into wells of a 6-well tissue culture plate. Precoat 

glass coverslip or delta-T dishes with matrix molecule of interest (e.g., 2 µg 

fibronectin/ml PBS) for 2 hr at 37 ◦ C. 

Other ECM molecules may be prepared in the same concentrations in PBS as fibronectin. 

Too high a ligand concentration will inhibit cell motility. If migration is slow or nonexistent, 

try decreasing the ligand concentration. 

Add cells 

2. Prepare a cell suspension of 1 × 10 6 70% confluent 24 hr serum-starved cells/ml in 

DMEM (see Basic Protocol 1, steps 4 to 5). 

Serum starve cells for 24 hr before initiating wound healing assay. 

Cells should be ∼70% confluent during starvation. 

3. Wash coverslips or delta-T dishes with PBS and plate 2 × 10 6 cells onto coverslips 

in a 6-well tissue culture plate or directly onto the delta-T dish. 

Coverslips in 6-well tissue culture plates are used if subsequent procedures include an 

imaging chamber. Bioptechs delta-T dishes are self-contained. 

4. Incubate at 37 ◦ C with 10% CO2 for 2 hr. 

The incubation time can be adjusted to accommodate other cells types if needed. This 

step is to facilitate equal cell adhesion and spreading on the matrix-coated glass surface. 


Wound the cultures 

5. Remove plates with coverslips from incubator. Using forceps to stabilize the coverslip 

in the well, carefully scratch the confluent monolayer of cells down the center using 

a micropipet tip. 

If using a delta-T dish, use a similar technique to scratch the center of the glass insert 

while holding the dish steady. 

This step is critical and will require practice in order to achieve consistent wounds. In 

order to reliably compare results from experiment to experiment, the wound distance must 

be equivalent each time. 

6. Rinse the coverslip/delta-T dish three times with 2 ml PBS to remove loose cells. 

If the cell monolayer is not washed effectively after scratching, loose cells can settle into 

the wounded area and result in an inaccurate interpretation of wound healing ability. 

7. Place coverslip in fresh DMEM medium with 0.5 µg/ml mitomycin-C and incubate 

at 37 ◦ C with 10% CO2 for 1 hr. 

Add 0.5 µg/ml mitomycin-C to all medium changes from this point. This will inhibit cell 

mitogenesis and, therefore, control for wound closure due to cell proliferation. 

Serum can be added (1% to 5% v/v, depending on cell type and required stimulation) at 

this point to stimulate cell motility if needed. 

Assess motility 

8. Aspirate medium and move coverslip into imaging chamber (if using the delta-T 

dishes, the imaging chamber is the unit itself). Add 1 ml of medium 199 with 

0.5 µg/ml mitomycin-C (with serum addition as needed). 

9. Carefully layer 0.5 ml mineral oil on top of the medium to prevent evaporation during 

imaging and place imaging chamber/delta-T dish in heated stage on the microscope 

and set to 37 ◦ C. 

10. Using a 10× or 20× objective, center the wound such that the image captured will 

show both sides of the cell monolayer flanking the wounded region. Using Improvision 

Openlab or comparable acquisition software, begin the time-lapse sequence 

and collect one image every 2 to 5 min for 12 to 15 hr or until the wound is closed. 

Alternatively, if time-lapse software is not available, etched-grid coverslips can be used 

(Bellco). These coverslips have a grid system that allows tracking and measuring wound 

closure over time with a light microscope while leaving the coverslips in the incubator in 

between phase-image collection. 

IMMUNOLOCALIZATION OF FOCAL ADHESION PROTEINS 

Immunostaining is the best approach to determining whether a protein of interest is 

localized to integrin-enriched focal adhesions. For valid interpretation of the staining 

results, two controls are essential: (1) immunoblot evaluation of the antibody against the 

protein before immunostaining and (2) cells costained with markers for focal adhesion 

proteins. The antibody to be used is suitable for cell staining analyses if it yields one 

band by immunoblotting analyses of whole cell lysates. The presence of multiple bands, 

even though the main one is the strongest, easily leads to misinterpretation of results. 

The most commonly used focal adhesion markers are vinculin and paxillin. Some of the 

best markers for activated focal adhesions are antibodies that recognize tyrosine-397phosphorylated 

FAK or active Src-family members. 

BASIC 

PROTOCOL 6 





14.7.21 


Analyzing FAK 



14.7.22 

Materials 

Matrix-coating substrates: prepared according to manufacturer’s directions and 

diluted (commonly to 10 µg/ml) in PBS (APPENDIX 2A) 

Fibronectin, human plasma (Roche) 

Laminin, human placenta (Sigma) 

Vitronectin, human plama (Sigma) 

Poly-L-lysine (70,000–150,000 or 150,000–300,000; Sigma) 

Collagen, Type I, human placenta (Calbiochem) 

Collagen, Type II, bovine (Calbiochem) 

Collagen, Type IV, human placenta (Calbiochem) 

Collagen, Type V, human (Calbiochem) 

FAK−/− and FAK+/+ cells (see Support Protocol 1) 

Replating and migration medium (see recipe), warm or growth medium (see 

recipe), 37◦C Phosphate-buffered saline (PBS; APPENDIX 2A) 

3.8% (w/v) paraformaldehyde fixative (see recipe) 

Acetone, cold (stored at −20◦C) or 

0.5% (v/v) Triton X-100/0.05% (v/v) Tween 20/PBS or 

0.2% Triton X-100/PBS or 

0.2% (v/v) Triton X-100/3.8% (w/v) paraformaldehyde/PBS or 

0.1% (v/v) Triton X-100/0.1% (w/v) sodium citrate or 

Methanol, cold 

Blocking antibody (e.g., ChromPure donkey IgG, unconjugated; Jackson 

Immunoresearch) or 

2% (w/v) BSA in PBS 

1to10µg/ml PBS (APPENDIX 2A) primary antibodies for focal adhesion markers 

anti-paxillin (ZO35; Zymed/Invitrogen) 

anti-vinculin (VIN-11-5; Sigma) 

anti-FAK (clone #77; BD-Transduction) 

anti-FAK (Ab-1; LabVision) 

anti-phospho Y397FAK (BioSource) 

anti-active Src family members (clone #28; BioSource) 

Secondary antibodies: e.g., fluorescein (FITC)-conjugated donkey antibodies 

(excitation/emission maxima 492/520 nm; Jackson Immunoresearch) or 

FITC-conjugate 

anti-mouse IgG 

anti-mouse IgM 

anti-rabbit IgG 

anti-goat IgG 

anti-rat IgG 

Rhodamine X (RRX)-conjugated donkey antibodies (excitation/emission maxima 

570/590 nm; Jackson Immunoresearch) or Rhodamine X (RRX)-conjugated 

anti-mouse IgG 

anti-mouse IgM 

anti-rabbit IgG 

anti-goat IgG 

anti-rat IgG 

Hoechst 33342 (excitation/emission maxima 350/460; Molecular Probes) 

Vectashield mounting medium (Vector) 

Nail polish, clear 

12-mm round coverslips, German glass (Bellco Glass) 

4-well tissue culture plates (Nunc) 

12-well tissue culture plates (Costar) 

18- to 22-G needle with a slightly bent tip 

Flat-ended forceps with beveled, unserrated tips, stainless steel (Millipore) 


6-well tissue culture plates 

Rotating platform shaker 

Vacuum source 

Porcelain spot plates with 12 cavities (CoorsTek) 

Light microscope with fluorescence excitation and detecting capability 

1. Coat 12-mm round coverslips with matrix-coating substrate (e.g., by incubating with 

10 µg/ml fibronectin 45 min at room temperature). 

The most commonly used adhesion molecule to induce formation of focal contacts is 

fibronectin, but the same volumes and concentrations are recommended for laminin, 

vitronectin, poly-L-lysine, and collagen. The coating time should be extended to 2 hr for 

collagen, and the coverslips should be washed with PBS to neutralize the acidity of the 

collagen solutions. 

Plate cells 

2. Prepare FAK−/− and FAK+/+ cell suspension of 2 × 10 4 cells/ml in 37 ◦ C replating 

and migration medium or growth medium (see Basic Protocol 1, steps 5 to 6), and 

add 0.5 ml cell suspension per well of a 4-well tissue culture plate containing a 

coated coverslip. 

Blocking with BSA is not required because cells will not attach to the uncoated glass 

surface. 

At this concentration the adherent cells will be spread apart. 

If the purpose is to determine activation or localization of the protein by adhesion molecule 

of interest, replating and migration medium should be used, otherwise growth medium is 

appropriate. 

The well size of 4-well tissue culture plates is the same as in 24-well plates. However, the 

wells are not so deep, and it is easier to take coverslips out. 

3. Incubate at 37 ◦ C for desired time. 

Activation (phosphorylation) of the focal adhesion proteins is the highest during cell 

spreading. For fibroblasts, the best time to capture localization of the activated proteins 

in focal adhesions is ∼1 hr after plating. At that time most of cells will be spread. Some 

cells will still be spreading and often both types can be visualized within a single field. 

Activated focal adhesion proteins at the leading edge of migrating cell are best visualized 

5 to 6 hr after wounding a cell monolayer with a micropipet tip. Activation is higher in 

the presence of serum. The most commonly used serum concentration is 10%, but one 

could also see activation at much lower serum concentrations. 

Fix and permeabilize cells 

4. Aspirate medium and rinse briefly 2 to 3 times in ∼1 ml PBS by adding ∼1mlPBS 

and aspirating or decanting it. 

For most cells 

5a. Add ∼1 ml 3.8% paraformaldehyde/PBS, pH 7.4 to fix cells. Incubate 20 min at 

room temperature and rinse with PBS as above. 

6a. Transfer coverslips into a porcelain spot plate filled with cold acetone (stored at 

−20 ◦ C) to permeabilize the cells. Keep on ice for 10 min. Remove acetone and add 

∼1 ml PBS to rehydrate for 3 to 5 min. 

Transfer of coverslips to a porcelain plate is necessary only when using acetone, which 

will dissolve plastic tissue culture dishes. 





14.7.23 


Analyzing FAK 



14.7.24 

Fixation and permeabilization are the most critical steps for successful immunostaining. 

Even for the same protein, this step can differ from cell type to cell type and from 

antibody to antibody. Most focal adhesion proteins are detectable when cells are fixed 

in paraformaldehyde and permeabilized in acetone. If the staining is weak or negative, 

changing fixation and permeabilization should be the first step to troubleshoot. 

For weakly staining cells (alternative 1) 

5b. Add ∼1 ml 3.8% (w/v) paraformaldehyde/PBS, pH 7.4 to fix cells. Incubate 20 min 

at room temperature. 

6b. Remove paraformaldehyde and add ∼1 ml 0.5% (v/v) Triton X-100 + 0.05% (v/v) 

Tween 20/PBS for 2 min to permeabilize the cells. 


5c. Add ∼1 ml 3.8% (w/v) paraformaldehyde/PBS, pH 7.4 to fix cells. Incubate 20 min 


6c. Remove paraformaldehyde and add ∼1 ml to 0.2% (v/v) Triton X-100/PBS for 2 

min to permeabilize the cells. 


5d. Add ∼1 ml 0.2% (v/v) Triton X-100/3.8% (w/v) paraformaldehyde/PBS and incubate 

2 min at room temperature to fix and permeabilize cells. Rinse several times with 

3.8% paraformaldehyde/PBS as in step 4. 

6d. Add ∼1 ml 3.8% paraformaldehyde/PBS and continue fixing for 20 min at room 

temperature. 


5e. Add ∼1 ml 3.8% (w/v) paraformaldehyde/PBS, pH 7.4 to fix cells. Incubate 20 min 


6e. Add ∼1 ml 0.1% (v/v) Triton X-100/0.1% (w/v) sodium citrate and keep 2 to 5 min 

on ice to permeabilize the cells. 


5f. Transfer coverslips into a porcelain spot plate filled with cold acetone (stored at 

−20◦C) to fix and permeabilize cells. Keep on ice for 10 min. 

In some cases in the more common protocol described in step 5a, paraformaldehyde 

fixation can interfere with staining. In these cases coverslips are directly exposed to 

acetone, which can also act as a fixative. 

6f. Remove acetone and add ∼1 ml PBS to rehydrate for 3 to 5 min. 

Staining of actin stress fibers is impossible when fixation/permeabilization is done with 

ONLY acetone or methanol (alternatives 5 and 6). 


5g. Add ice-cold methanol (stored at −20◦C) to fix and permeabilize cells. Keep on ice 

for 10 min. 

6g. Aspirate methanol, rinse briefly in ∼1 ml PBS and leave in PBS for 3 to 5 min to 

rehydrate. 

Staining of actin stress fibers is impossible when fixation/permeabilization is done with 

ONLY acetone or methanol (alternatives 5 and 6). 


Block and immunostain cells 

7. Pace a 30-µl drop of PBS containing blocking antibodies and primary antibodies for 

focal adhesion markers on the inner side of 12-well plate lid (facing up), and place 

the coverslip with fixed and permeabilized cells on top, with the attached cells facing 

the drop of antibody solution. Cover with the bottom of the plate and incubate 1 hr 

at room temperature or as long as overnight at 4 ◦ C, depending on convenience and 

the effectiveness of staining with various samples and procedures. 

Using a 30-µl drop of PBS containing blocking antibodies and primary antibodies for 

focal adhesion markers on the inner side of a 12-well plate lid helps prevent evaporation, 

ensure equal exposure, and use less antibody solution. 

Flip coverslip over the 30-µl drop of PBS containing blocking and primary antibodies 

using an 18- to 22-G needle with a slightly bent tip to lift and flat-ended forceps to hold 

and transfer the coverslip. Sharp-ended forceps tend to crack the coverslip. Alternatively, 

if the cell type is sensitive to flipping the coverslips for incubations and cells detach, 

utilize a humidified chamber (to avoid evaporation) and add 50 µl of antibody directly 

on top of the coverslips. 

Blocking is done with whole IgG of the host animal for the secondary antibody. For 

example, the primary antibody is made in mouse. If the secondary antibody is fluoresceinconjugated 

donkey anti-mouse, donkey whole IgG should be used for blocking. If the 

secondary antibody is fluorescein-conjugated goat anti-mouse, goat whole IgG should 

be used for blocking, etc. Blocking antibodies are sold in concentrations ∼10 mg/ml. 

Recommended dilution for blocking purposes is 1:100 (∼100 µg/ml). To keep costs low, 

the authors use secondary antibodies that are all made in donkey, requiring only one 

reagent (ChromPure donkey whole IgG) for blocking purposes. 

Alternatively, blocking can be performed with 2% (w/v) BSA in PBS. In this case, blocking 

is done at the same time as sample incubation with the primary antibody, saving time. BSA 

blocking is less specific than IgG, and in rare cases BSA might be too strong a blocking 

agent. 

Dilution of primary antibodies can vary and should be optimized for each antibody. 

Generally a range of 1 to 10 µg/ml is effective. A starting point for optimization would 

be a 1:100 dilution of a commercial antibody, which is usually sold in concentrations 0.1 

to1mg/ml. 

For co-immunostaining, incubate samples with both primary antibodies and with blocking 

antibodies at the same time. If incubations are for several hours, samples can be left at 

room temperature. If incubation is performed overnight, the incubation should be at 4◦C. 8. Lift the coverslip by injecting 150 µl PBS under it, and place it in a well of a 6-well 

tissue culture plate, attached cells facing up. Wash in three changes of ∼3 mlPBS 

5 min each time at ∼50 rpm on rotating platform shaker. 

Smaller size wells (e.g., wells of 12- or 24-well plates) do not allow sufficient movement 

of PBS to wash unbound antibodies from the samples. 

9. Pace a 30-µl drop of PBS containing secondary antibodies and 10 µg/ml Hoechst 

33342 on the inner side of a 12-well plate lid (facing up), and place the coverslip 

with fixed and permeabilized cells on top with the attached cells facing the drop of 

antibody solution. Cover with plate bottom and incubate 40 min at room temperature. 

Hoechst 33342 binds DNA, and it is an excellent bright nuclear marker visible at UV 

excitation wavelengths. It does not leak to green or red channels. It is of immense help 

for orientation and focusing of samples during microscopy. 

For visualization of actin stress fibers, fluorescent phalloidin conjugate is added together 

with secondary antibody for costaining of two antigens: one is detected by the primary 

antibody, and the other (actin stress fibers) is detected directly by fluorophore-conjugated 

phalloidin. 





14.7.25 


ALTERNATE 

PROTOCOL 4 

Analyzing FAK 



14.7.26 

10. Wash in three changes of ∼3 ml PBS three times 5 min each time at ∼50rpmona 

rotating platform shaker. 

11. Mount on microscope slides in Vectashield mounting medium. Aspirate excess 

mounting medium under the coverslip using a Pasteur pipet attached to vacuum. 

To prevent sliding, fix the edge of the coverslip with clear nail polish. Wait 15 to 20 

min for the nail polish to dry, then analyze samples under the microscope. 

Samples can be preserved at 4 ◦ C in the dark for varying lengths of time, depending on 

the strength of the staining. For example, actin staining can be preserved for a month 

or longer. Weak staining will fade beyond recognition within several days. The authors 

recommend analysis within 2 to 3 days after staining. 

IMMUNOLOCALIZATION OF ACTIN STRESS FIBERS 

Visualization of actin stress fibers with fluorescent phalloidin, a toxin with high binding 

affinity for actin filaments, is easy and almost certainly yields dazzling results. Unlike 

indirect immunostaining with antibodies, phalloidin staining is strong, and the contrast 

between stained and unstained areas is high. Because focal adhesions are located at 

the end of actin stress fibers, phalloidin staining is also used as a localization marker 

and as a quick assay to assess cell shape. Costaining actin stress fibers with fluorescent 

phalloidin–conjugate and DNA-binding Hoechst dye can be a rewarding experience for 

a new investigator who is performing immunofluorescence cell staining for the first time. 


0.4 to 1 U/ml (10 to 30 nM) fluorescein-conjugated phalloidin (Molecular Probes) 

or rhodamine-conjugated phalloidin (Molecular Probes) in PBS 

Rotating platform shaker 

Vacuum source 

Prepare cells 

1. Coat coverslips, prepare FAK−/− and FAK+/+ cell suspension, and plate the cells 

(Basic Protocol 6, steps 1 to 3). 

2. Incubate at 37 ◦ C for desired time. 

Fix and permeabilize cells 

3. Aspirate medium and rinse briefly 2 to 3 times in ∼1 ml PBS by adding ∼1mlPBS 

and aspirating or decanting it. 

4. Add ∼1 ml 3.8% paraformaldehyde/PBS, pH 7.4, to fix cells. Incubate 20 min at 

room temperature and rinse with PBS as above. 

5. Transfer coverslips into a porcelain spot plate filled with cold acetone (stored at 

−20 ◦ C) to permeabilize the cells. Keep on ice for 10 min. Remove acetone and add 

∼1 ml PBS to rehydrate for 3 to 5 min. 

6. Place a 30-µl drop containing 0.4 to 1 U/ml (10 to 30 nM) fluorescein-conjugated 

phalloidin with 10 µg/ml Hoechst 33342 in PBS on the inner side of a 12-well tissue 

culture plate lid (facing up), and place the coverslip with fixed and permeabilized 

cells on top with the attached cells facing the drop of solution. Cover with the plate 

bottom and incubate 30 min at room temperature. 

If dissolved as recommended by the manufacturer (Molecular Probes), 0.4 to 1 U/ml (10 

to 30 nM) phalloidin is 1:200 to 1:500. 


Rhodamine-conjugated phalloidin is an alternative to fluorescein-conjugated phalloidin. 

If costaining with some other protein, phalloidin should be added together with the 

secondary antibody (see Basic Protocol 6). 

7. Wash in three changes of ∼3 ml PBS 5 min each time at ∼50 rpm on a rotating 

platform shaker. 

8. Mount on microscope slides in Vectashield mounting medium. Aspirate excess 

mounting medium under the coverslip using a Pasteur pipet attached to vacuum. 

To prevent sliding, fix the edge of the coverslip with nail polish. Wait 15 to 20 min 

for the nail polish to dry, then analyze samples under the microscope. 

Samples can be preserved at 4 ◦ C in the dark for varying lengths of time depending on 

the strength of the staining. For example, actin staining can be preserved for a month 

or longer. Weak staining will fade beyond recognition within several days. The authors 

recommend analysis within 2 to 3 days after staining. 


Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see 


BSA blocking buffer 

20 mM Tris·Cl, pH 7.4 

150 mM NaCl 

2% (w/v) BSA (Fraction V) 

0.05% (v/v) Tween 20 

0.05% (w/v) sodium azide 

Store up to 3 months at 4 ◦ C 

Cell growth medium 

DMEM high-glucose with L-glutamine (Invitrogen) 

10% (v/v) FBS 

1 mM sodium pyruvate (Invitrogen) 

0.1 mM MEM nonessential amino acids (Invitrogen) 

10 U/ml penicillin and 10 µg/ml streptomycin (Invitrogen; 100× stock) 

Sterilize by passing through a 0.22-µm filter 

Store up to 2 weeks at 4 ◦ C 

Coomassie blue stain 

25% (v/v) isopropanol (2-propanol) 

10% (v/v) glacial acetic acid 

0.03% (w/v) Coomassie G-250 (BioRad) 

Store up to 1 year at 21 ◦ C 

Destaining solution 

10% (v/v) isopropanol (2-propanol) 

10% (v/v) glacial acetic acid 

Store up to 1 year at 21 ◦ C 

FAK-Pyk2 kinase buffer 



10 mM MgCl2 

10 mM MnCl2 

150 mM NaCl 






14.7.27 


Analyzing FAK 



14.7.28 

HNTG buffer 


0.1% (v/v) Triton X-100 

150 mM NaCl 



Lac Z staining solution 

70 ml H2O 

10 ml of 10× PBS (1× final) 

10 ml of 50 mM potassium ferricyanide (5 mM final) 

10 ml of 50 mM potassium ferrocyanide (5 mM final) 

0.2mlof1mlMgCl2 (2 mM final) 

Store up to 2 weeks at 4 ◦ C 

Just before use, add 1 ml of 20 mg/ml Xgal per 20 ml of staining solution (1 mg/ml 

final). Make a stock solution of 20 mg/ml X-gal in dimethylformamide (DMF). 

Store at −20 ◦ C in the dark. 

NOTE: Use a glass pipet to measure DMF solutions as it will dissolve plastics. 

Laemmli SDS sample buffer (2×) 



4% (w/v) SDS 

0.006% (w/v) bromophenol blue (Fisher Biotech) 

3.5% (v/v) 2-mercaptoethanol 

Store up to 1 week at 4 ◦ C or 6 months at −20 ◦ C 

Magnesium/ATP cocktail 

75 mM MgCl2 

500 µM ATP 

20 mM MOPS, pH 7.2 

25 mM β-glycerol phosphate 

5mMEGTA 

1 mM sodium orthovanadate 

1 mM dithiothreitol 

Store up to 1 week at 4 ◦ C 

Paraformaldehyde fixative, 3.8% (w/v) 

Add 3.8 g paraformaldehyde (Sigma) to 100 ml PBS (APPENDIX 2A), heat, and stir to 

dissolve. When temperature reaches 50 ◦ C, add ∼100 µl 1 N NaOH, and continue 

stirring until temperature reaches 60 ◦ C. Remove from the heating plate, continue 

stirring until it reaches room temperature and filter to remove fine precipitate. Store 

up to 10 days at 21 ◦ C. 

Once made, paraformaldehyde fixative is ready to use for up to 10 days. The exact 

percentage of paraformaldehyde is not crucial; it is sufficient to be somewhere 

between 3.8% and 4.0%. Similarly, the exact pH does not matter, if it is somewhere in 

a physiological range (7.2 to 7.6). Paraformaldehyde from different manufacturers 

or of different purity might require different amount of NaOH to adjust the pH. 

Replating and migration medium 

500 ml DMEM 

0.5% (w/v) BSA (add 20 ml 12.5% BSA solution) 

1 mM sodium pyruvate (Invitrogen) 

continued 


0.1 mM MEM nonessential amino acids (Invitrogen) 


Store up to 1 month at 4 ◦ C 

RIPA cell lysis buffer 


1% (v/v) sodium deoxycholate 

0.1% (w/v) SDS 


150 mM NaCl 


1.5 mM MgCl2 

1mMEGTA 

10 mM sodium pyrophosphate 

100 mM sodium fluoride 

1 mM sodium orthovanadate (add dropwise) 





Add ingredients slowly to water. Start over if buffer becomes brown after sodium 

orthovanadate addition. 

Alternatively, a protease inhibitor cocktail (e.g. Sigma P2714) may be used instead 

of aprotinin and leupeptin. 

Src kinase buffer 

20 mM PIPES, pH 7.0 

10 mM MnCl2 

1mMDTT 


Tris-buffered saline with Tween (TBST) 


150 mM NaCl 

0.05% (v/v) Tween 20 


Triton lysis buffer 

RIPA cell lysis buffer (see recipe) without sodium deoxycholate and SDS addition. 


Trypsin inhibitor solution 

500 ml DMEM 

125 mg soybean trypsin inhibitor (Worthington) 

Dissolve by stirring 

0.25% (w/v) BSA (add 10 ml of 12.5% w/v BSA solution) 

Sterilize by passing through a 0.22 µm filter 

Prepare fresh 

Western blot stripping buffer 


2% (w/v) SDS 

100 mM 2-mercaptoethanol 






14.7.29 


Analyzing FAK 



14.7.30 

COMMENTARY 


FAK was first identified in 1992 as a 

highly tyrosine-phosphorylated protein associated 

with cellular focal adhesions, i.e., the 

integrin-associated cell attachment points with 

the extracellular matrix (Parsons, 2003). At 

that time, it was shown that changes in cell 

adhesion or integrin clustering could result 

in the increased phosphorylation of tyrosine 

in FAK (Guan and Shalloway, 1992; Hanks 

et al., 1992; Kornberg et al., 1992; Schaller 

et al., 1992). Today we know that this activation 

event is much more complicated than the 

simple binding of FAK to integrins leading 

to intermolecular FAK phosphorylation after 

clustering (Schlaepfer and Mitra, 2004). For 

instance, evidence to date does not strongly 

support direct binding of FAK to integrins. 

Instead, FAK is associated with integrin cytoplasmic 

tails through binding to integrinassociated 

proteins such as paxillin and talin. 

Additionally, integrins can activate Src-family 

protein-tyrosine kinases in a manner that is 

independent of FAK. It is known that cell 

binding to fibronectin promotes FAK and 

c-Src activation and the formation of a 

FAK-Src signaling complex, and these facts 

support the conclusion that the events are related. 

Current research is focused on deciphering 

the molecular mechanisms through which 

FAK and c-Src are activated by integrins, and 

the methods outlined in this unit comprise the 

foundation for these studies. 

Additional complexity in this integrin signaling 

linkage comes from the studies of the 

FAK-related kinase, Pyk2, which is highly expressed 

in FAK−/− cells (Sieg et al., 1998). 

Stimulation of FAK−/− cells by replating on 

fibronectin promotes the activation of Srcfamily 

tyrosine kinases and the enhanced tyrosine 

phosphorylation of Pyk2. However, neither 

Pyk2 nor c-Src strongly colocalize with 

FAK−/− cell focal adhesions, and these cells 

do not correctly spread and migrate like wildtype 

FAK+/+ fibroblasts (Sieg et al., 1999; 

Klingbeil et al., 2001). The localization of 

FAK to particular integrin signaling sites is important 

for fibronectin-stimulated cell motility, 

the molecular mechanisms of which are still 

under investigation. 

What the authors have tried to accomplish 

in this unit is to consolidate related protocols 

on how to measure changes in FAK, c-Src, and 

Pyk2 activity after fibronectin-mediated integrin 

stimulation through the use of in vitro kinase 

assays, phospho-specific blotting, or im- 

munolocalization techniques. These assays involve 

using FAK−/− and FAK+/+ cells. The 

handling or preparation of these cells prior to 

performing biochemical activation assays is 

the same as the procedures used in evaluating 

the properties of these cells in cell migration 

assays. In this manner, cause and effect relationships 

can be established between integrinstimulated 

signaling events and the modulation 

of a complex biological process such 

as cell motility. Alternate protocols that have 

been successfully used by other investigators 

include the use of antibodies or recombinant 

ligands for particular integrin subunits to facilitate 

binding or clustering and intracellular 

signaling (Giancotti and Ruoslahti, 1999). 

As cells usually contain multiple integrin α/β 

pairs that can bind fibronectin, and current research 

has shown that sequence-specific differences 

between integrin cytoplasmic domains 

facilitates the binding or activation of distinct 

intracellular signaling proteins (Liu et al., 

1999; de Virgilio et al., 2004), antibody/ligand 

activation of integrins is a powerful means to 

study integrin signaling. However, antibodymediated 

clustering of integrins is usually a 

weaker stimulus compared to cell binding and 

spreading on matrix proteins. Therefore, if a 

recombinant ligand can be purified and immobilized 

to facilitate specific integrin engagement 

and cell binding, this can be effectively 

used to study the signaling linkage between 

particular integrins, the actin cytoskeleton, and 

cell motility. 



Maintenance of FAK+/+ AND FAK−/− 

fibroblasts and replating assays for 

signaling studies 

It is critical to use well maintained 

FAK+/+ and FAK−/− cells for all experiments 

in this unit. Because the fibroblasts 

may exhibit altered morphology and premature 

senescence when overly confluent or at 

high passage number, cells need to be subcultured 

consistently and used within the first 

15 passages from the cells available from 

American Type Culture Collection (ATCC). 

To prevent poor attachment to the extracellular 

matrices when cells are replated, avoid overtrypsinization 

and always make fresh trypsin 

inhibitor solution. Further, serum starvation 

is an important step in allowing maximal 

cell response and activation of focal adhesion 

molecules to stimuli (e.g., fibronectin). This 


is also critical for successful motility assays 

(Basic Protocol 4) and scratch-wound healing 

assays (Basic Protocol 5). 

In vitro kinase assay 

It is important that a control antibody is included 

with immunoprecipitation experiments 

to accurately interpret in vitro kinase assay 

results. For example, preimmune serum can 

serve as the control for a rabbit antiserum. 

Further, the species (e.g., rabbit, mouse, goat, 

rat, or sheep) of antibody used in immunoprecipitation 

should be carefully considered if 

the immunoprecipitates will be analyzed by 

subsequent immunoblotting. HRP-conjugated 

secondary antibodies will recognize the immunoprecipitating 

antibody as well as the primary 

immunoblotting antibody of the same 

species, resulting in very high background 

around the 60-kDa protein region. As this is 

region where c-Src is present on gels, nonspecific 

antibody cross-reactivity can complicate 

co-immunoprecipitation experiments and 

c-Src immunodetection. 

Immunoblotting with FAK/Pyk2 

phospho-specific antibodies 

For the detection of FAK and Pyk2 phosphorylation 

with phospho-specific antibodies, 

the most critical issue is the specificity and 

quality of the antibody. Ideally, the phosphospecific 

antibody should recognize only the 

phosphorylated form of the target protein on 

a specific site. However, in the experience of 

the authors, some phospho-specific antibodies 

for FAK can cross-react with corresponding 

sites on Pyk2 and may even react with other 

phosphorylation sites on the target protein. 

It is for this reason that results should be 

double-checked using complementary means 

such as phospho-specific antibodies from various 

manufacturers and alternate procedures 

(e.g., in vitro kinase assays). When reprobing 

with different phospho-specific antibodies, it 

is critical to ensure that stripping or removal 

of previous antibodies is complete. This can 

be done by performing the ECL reaction and 

exposing film after every blot stripping to confirm 

that no residual signal remains. 

Haptotaxis motility assays 

As confluent cells do not respond to extracellular 

matrix stimuli, it is essential to maintain 

a low cell confluency when performing the 

haptotaxis assay. Additionally, chamber coating 

is key to a successful and interpretable 

motility assay. Uneven or improper chamber 

coating will result in experimental variability 

within single chambers as well as through- 

out the experiment. Potential causes of uneven 

coating include denatured or sticky extracellular 

proteins and the presence of bubbles underneath 

the chambers during coating. 

Scratch-wound healing assays 

After generating the wound, it is important 

to ensure that no residual cells settle in the 

scratch area. This would severely complicate 

interpretation of the assay and can be avoided 

by thorough washing after creating the scratch 

wound. Additionally, wound-closure measurements 

must be compared to the original wound 

distance for each individual coverslip due to 

experimental scratch variations. 

Immunostaining 

The most important issues for visualization 

of focal adhesion molecules with immunofluorescent 

staining are (1) fixation and permeabilization 

of cells and (2) specificity of the 

staining antibody. Optimized fixation allows 

the preservation and localization of the antigen 

as well as cell morphology, while proper 

permeabilization allows access to the antigen. 

To achieve specificity for the immunostaining, 

the primary antibody should recognize 

only its target antigen, and proper controls 

(e.g., preimmune serum or secondary antibody 

only) should be included to optimize the staining 

conditions and to reduce background. See 

Table 14.7.3 for troubleshooting the experiments 

described in this unit. 

The authors do not recommend use of 

preimmune serum as a control for staining 

specificity. The more appropriate control is a 

Western blot of whole cell lysate made from 

the same cells treated in the same matter. If 

multiple bands are visible, the antibody is not 

reliable for immunostaining. A control for secondary 

antibody is not necessary if whole IgG 

of the same species in which the secondary 

antibody was generated is used as a blocking 

agent together with the primary antibody. 


In attached and serum-starved FAK+/+ 

and FAK−/− cells, FAK and Pyk2, respectively, 

display high basal levels of tyrosine 

phosphorylation and moderate level of kinase 

activity. In suspended cells, phosphorylation 

of tyrosine in both FAK and Pyk2 

is greatly reduced. Replating cells on fibronectin, 

but not poly-L-lysine, increases tyrosine 

phosphorylation on both FAK and Pyk2 

in FAK+/+ and FAK−/− cells. Immunoblotting 

with phospho-specific antibodies reveals 

that FAK and Pyk2 can be phosphorylated on 





14.7.31 


Analyzing FAK 



14.7.32 

Table 14.7.3 Troubleshooting Guide for Common Problems Encountered in These Assays 

Problem Possible cause Solution 

Cell replating 

Cells slow to adhere 

and spread on matrix 

In vitro kinase assay 

No or low levels of 

substrate labeling 

Too much trypsin used to 

prepare cell suspension 

Use limited trypsin/EDTA treatment at 

37◦C and tap plates to remove cells 

Make fresh trypsin inhibitor solution 

Old [γ- 32 P]ATP Order fresh [γ- 32 P]ATP 

Kinase denatured or 

degraded 

High background Carryover of cell debris 

following preclearance 

Phospho-specific 

immunoblotting 

Phospho-specific 

antibody recognizing 

nonphosphorylated 

form of protein in 

immunoblotting 

Haptotaxis motility 

assay 

Migratory cells too 

dense to quantify 

Inconsistent cell 

migration 

Scratch-wound 

healing 

Cells do not close the 

wound 

Cells close the wound 

too quickly 

Add fresh protease inhibitor to lysis 

buffer. Use freshly prepared lysates for 

immunoprecipitation 

Avoid repeated freeze-thaw of samples 

Use polyclonal antibodies to target 

protein for immunoprecipitation 

Avoid disturbing cell debris when 

transferring supernatant during lysate 

preparation 

Incomplete washing Completely resuspend beads at each 

wash 

Too much antigen or 

antibody present 

Optimize the amount of lysates and 

concentration of antibody used. Reduce 

antibody incubation time and increase 

washing 

Assay went too long Decrease assay time 

Too many cells were plated Plate fewer cells 

Improper chamber coating Avoid bubbles underneath chamber 

Matrix protein precipitated 

out of solution 

Matrix protein 

concentration is too high 

Cells are not plated as a 

confluent monolayer 

Cells require serum 

stimulation 

Confirm the pH of medium used to 

resuspend matrix protein 

Decrease matrix protein concentration 

Increase cell density 

Add 1% to 5% serum during assay 

Too many cells Decrease cell density 

Elevated cell proliferation Add mitomycin C to block cell division 

continued 


Table 14.7.3 Troubleshooting Guide for Common Problems Encountered in These Assays, 

continued 


Immunolocalization 

No staining Primary antibody is not 

binding to epitope 

Staining looks 

non-specific or high 

background 

Identical pattern 

observed for multiple 

proteins 

Signal “leaking” 

through microscope 

filters 

Complications due to 

fixation procedure 

Primary antibody does not 

cross-react across species 

Antigen target is expressed 

at low levels 

Primary antibodies are 

from same species 

Too high concentration of 

primary or secondary 

antibody 

multiple tyrosine residues following replating 

onto fibronectin matrix, and, following integrin 

activation, Src-associated in vitro kinase 

assays demonstrate that FAK and Pyk2 can 

transiently form signaling complexes with Src. 

In haptotaxis and scratch-wound healing 

assays, FAK−/− cells display impaired migration 

compared to FAK+/+ cells. The defective 

cell motility of FAK−/− cells cannot 

be compensated by the presence of high 

levels of Pyk2, but it can be rescued by transiently 

re-expressing FAK. Furthermore, FAK 

can be detected via immunofluorescence staining 

that prominently and specifically localize 

to focal adhesions. In fact, some of the best 

markers of activated focal adhesions are antibodies 

that recognize FAK phosphorylated 

on tyrosine 397. Finally, strong activation of 

FAK can be detected in focal adhesions about 

1 hr after replating fibroblasts onto extracellular 

matrix as well as at the leading edge of 

migration 5 to 6 hr following a scratch wound. 


Cells need to be serum starved (Support 

Protocol 2) for 16 to 24 hr before replating 

(Basic Protocol 1), haptotaxis motility (Basic 

Use different primary antibody 

Try another fixation procedure 

Try a different secondary antibody 

Try another fixation procedure 

Change primary antibody 

Increase primary antibody incubation 

from 1 hr to overnight. 

Increase concentration of primary 

antibody 

Use biotin-avidin enhancement 

Change one primary antibody to a 

different species 

Decrease antibody concentrations or 

increase number of washes 

Protocol 4), and scratch-wound healing 

(Basic Protocol 5) assays. Cells are kept in 

suspension for at least 45 min or replated 

for 1 to 3 hr before whole-cell lysates are 

collected, which takes 1 hr to complete 

(Support Protocol 3). 

FAK, Pyk2, and c-Src can be captured from 

whole cell lysates by immunoprecipitation in 4 

to 5 hr (Support Protocol 4), and can be further 

analyzed through in vitro kinase assay (Basic 

Protocol 2) or processed by immunoblotting 

for their phosphorylation and activation status 

(Basic Protocol 3). In these protocols SDS- 

PAGE gel electrophoresis takes 1 to 1.5 hr, 

gel transfer onto membrane takes 1.5 to 2 hr, 

and immunoblotting is best when membranes 

are incubated in primary antibodies overnight. 

Therefore, these biochemical assays will take 

a total of 3 days. 

Following serum starvation (16 to 24 hr), 

the haptotaxis assay (Basic Protocol 4) takes 

∼8 hr to complete. It takes 2.5 to 3 hr to prepare 

the chambers and 1 to 1.5 hr to prepare cells 

(depending on how many different experimental 

groups are being analyzed), but these two 

items can be done concurrently. Cells migrate 

for 3 hr followed by an additional 1 to 2 hr 





14.7.33 


Analyzing FAK 



14.7.34 

to fix and stain chambers. Microscopic quantification 

of migratory cells can take several 

hours. 

To transfect FAK+/+ and FAK−/− cells 

(Alternate Protocol 3), cells need to be plated 

the previous day; preparation of transfection 

mix requires 45 min to 1 hr, and the transfection 

takes 5 hr. Cells are grown for an additional 

48 hr to allow maximal protein expression 

and need to be serum starved for 24 hr 

before haptotaxis assay. 

For the scratch-wound healing assay (Basic 

Protocol 5), cells are serum starved for 

16 to 24 hr, plated on a cover slip for 2 

hr, scratch wounded, and incubated for 1 hr 

after wounding. Live images are collected for 

12 to 15 hr. The scratch-wound healing assay 

spans 32 to 45 hrs, but requires only 2 to 3 hr 

of hands-on time. For immunolocalization of 

focal adhesion proteins (Basic Protocol 6), immunofluorescence 

staining can be performed 

in ∼7hr. 

Internet Resources 

http://www.cellmigration.org/index.shtml 

This cell migration gateway represents a unique 

collaboration between the Cell Migration Consortium 

(CMC) and Nature Publishing Group (NPG) 

and is designed to facilitate navigation through the 

complex world of cell migration research. 

http://www.signaling-gateway.org 

This signaling gateway represents a unique collaboration 

between academia and scientific publishing 

and is designed to facilitate navigation of the complex 

world of research into cellular signaling. Information 

and data presented here are freely available 

to all. 


de Virgilio, M., Kiosses, W.B., and Shattil, S.J. 

2004. Proximal, selective, and dynamic interactions 

between integrin αIIbβ3 and protein 

tyrosine kinases in living cells. J. Cell Biol. 

165:305-311. 

Engebrecht, J., Brent, R., and Kaderbhai, M.A. 

1991. Minipreps of plasmid DNA. In Current 

Protocols in Molecular Biology (F.M. Ausubel, 

R. Brent, R.E. Kingston, D.D. Moore, 

J.G. Seidman, J.A. Smith, and K. Struhl, eds.) 

pp. 1.6.1-1.6.10. John Wiley & Sons, Hoboken, 

N.J. 

Giancotti, F.G. and Ruoslahti, E. 1999. Integrin signaling. 

Science 285:1028-1032. 

Guan, J.L. and Shalloway, D. 1992. Regulation of 

focal adhesion-associated protein tyrosine kinase 

by both cellular adhesion and oncogenic 

transformation. Nature 358:690-692. 

Hanks, S.K., Calalb, M.B., Harper, M.C., and 

Patel, S.K. 1992. Focal adhesion protein- 

tyrosine kinase phosphorylated in response to 

cell attachment to fibronectin. Proc. Natl. Acad. 

Sci. U.S.A. 89:8487-8491. 

Hynes, R.O. 2002. Integrins: Bidirectional, 

allosteric signaling machines. Cell 110:673- 

687. 

Ilic, D., Furuta, Y., Kanazawa, S., Takeda, N., 

Sobue, K., Natkatsuji, N., Nomura, S., Fujimoto, 

J., Okada, M., Yamamoto, T., and Aizawa, S. 

1995. Reduced cell motility and enhanced focal 

adhesion contact formation in cells from fakdeficient 

mice. Nature 377:539-544. 

Klingbeil, C.K., Hauck, C.R., Hsia, D.A., Jones, 

K.C., Reider, S.R., and Schlaepfer, D.D. 2001. 

Targeting Pyk2 to β1-integrin-containing focal 

contacts rescues fibronectin-stimulated signaling 

and haptotactic motility defects of 

focal adhesion kinase-null cells. J. Cell Biol. 

152:97-110. 

Kornberg, L., Earp, H.S., Parsons, J.T., Schaller, 

M., and Juliano, R.I. 1992. Cell adhesion or 

integrin clustering increases phosphorylation of 

a focal adhesion-associated tyrosine kinase. J. 

Biol. Chem. 267:23439-23442. 

Liu, S., Thomas, S.M., Woodside, D.G., Rose, 

D.M., Kiosses, W.B., Pfaff, M., and Ginsberg, 

M.H. 1999. Binding of paxillin to α4 

integrins modifies integrin-dependent biological 

responses. Nature 402:676-681. 

Parsons, J.T. 2003. Focal adhesion kinase: The first 

ten years. J. Cell Sci. 116:1409-1416. 

Schaller, M.D., Borgman, C.A., Cobb, B.A., 

Vines, R.R., Reynolds, A.B., and Parsons, J.T. 

1992. pp125fak a structurally distinctive proteintyrosine 

kinase associated with focal adhesions. 

Proc. Natl. Acad. Sci. U.S.A. 89:5192-5196. 

Schlaepfer, D.D. and Mitra, S.K., 2004. Multiple 

connections link FAK to cell motility and invasion. 

Curr. Opin. Genet. Dev. 14:92-101. 

Schlaepfer, D.D., Hauck, C.R., and Sieg, D.J. 1999. 

Signaling through focal adhesion kinase. Prog. 

Biophys. Mol. Biol. 71:435-478. 

Schwartz, M.A. 2001. Integrin signaling revisited. 

Trends Cell Biol. 11:466-70. 

Seidman, C.E., Struhl, K., Sheen, J., and Jessen, T. 

1997. Introduction of plasmid DNA into cells. 

In Current Protocols in Molecular Biology (F.M. 

Ausubel, R Brent, R.E. Kingston, D.D. Moore, 

J.G. Seidman, J.A. Smith, and K. Struhl, eds.) 

pp. 1.8.1-1.8.10. John Wiley & Sons, Hoboken, 

N.J. 

Sieg, D.J., Ilic, D., Jones, K.C., Damsky, C.H., 

Hunter, T., and Schlaepfer, D.D. 1998. Pyk2 and 

Src-family protein-tyrosine kinases compensate 

for the loss of FAK in fibronectin-stimulated 

signaling events but Pyk2 does not fully function 

to enhance FAK- cell migration. EMBO J. 

17:5933-5947. 

Sieg, D.J., Hauck, C.R., and Schlaepfer, D.D. 1999. 

Required role of focal adhesion kinase (FAK) for 

integrin-stimulated cell migration. J. Cell Sci. 

112:2677-2691. 


Contributed by Joie A. Bernard-Trifilo, 

Ssang-Taek Lim, Shihe Hou, and 

David D. Schlaepfer 

The Scripps Research Institute 

La Jolla, California 

Dusko Ilic 

Stem Life Line, Inc. 

San Carlos, California 





14.7.35 


Rho GTPase Activation Assays 

Stéphanie Pellegrin 1 and Harry Mellor 1 

1 Mammalian Cell Biology Laboratory, Department of Biochemistry, School of Medical 

Sciences, University of Bristol, United Kingdom 

ABSTRACT 

The Rho GTPase family of signaling proteins controls a wide range of highly dynamic 

cellular processes. Activation of Rho GTPases can be investigated and quantified in cell 

extracts using so-called pull-down assays. Proteins that bind specifically to the activated 

form of the Rho GTPase are used to capture it onto a bead support. Western blotting of 

the captured samples with specific antibodies then allows for quantification of the level 

of Rho GTPase activation in the sample. This unit describes the techniques for preparing 

the reagents required for assays of RhoA, Rac, and Cdc42 and gives practical tips for 

the successful application of the assay in a range of situations. Curr. Protoc. Cell Biol. 

38:14.8.1-14.8.19. C○ 2008 by John Wiley & Sons, Inc. 

Keywords: Rho GTPase � cytoskeleton � cell signaling 

INTRODUCTION 

Most Rho GTPases cycle between an active GTP-bound and an inactive GDP-bound 

state. When bound to GTP, Rho GTPases can interact with their effector proteins, and 

this nucleotide-dependent interaction forms the basis of the activation assays described 

below. The Rho GTPase-binding domain (RBD) of Rho GTPase effectors can be made 

Figure 14.8.1 Diagram of Rho-binding domains used for GST pull-down assays: (A) Rhotekin- 

RBD and (B) PAK-CRIB. The GST-RBD constructs we have used span residues 7 to 89 of Rhotekin 

variant 2 (accession number NM 033046) and residues 1 to 253 of PAK1 (accession number 

NM 002576). When made as GST-fusion proteins, these are 35.7-kDa and 55.7-kDa respectively. 

Abbreviations: HR1, protein kinase C-related kinase homology region 1; PH, pleckstrin homology 

domain; RBD, Rho binding domain. 

Current Protocols in Cell Biology 14.8.1-14.8.19, March 2008 

Published online March 2008 in Wiley Interscience (www.interscience.wiley.com). 

DOI: 10.1002/0471143030.cb1408s38 


UNIT 14.8 



14.8.1 

Supplement 38

BASIC 

PROTOCOL 

Rho GTPase 

Activation Assays 

14.8.2 

as a glutathione-S-transferase (GST) fusion protein, coupled to beads, and added to cell 

lysates to pull down the active, GTP-bound form of the Rho GTPase (see UNIT 17.5). The 

RhoA binding domain from Rhotekin has been widely used to pull down active RhoA 

(Ren et al., 1999; Ren and Schwartz, 2000), whereas the PAK-CRIB domain of PAK1 is 

favored for Rac1 and Cdc42 activation assays (Sander et al., 1998; Benard et al., 1999; 

Benard and Bokoch, 2002). These constructs are shown in Figure 14.8.1. The CRIB 

motif of WASP has also been used for Cdc42 activation assays (Haddad et al., 2001). 

This unit describes how to make recombinant GST-PAK-CRIB beads for Rac1 or Cdc42 

activation assays and recombinant GST-Rhotekin-RBD beads for RhoA activation 

assays (Support Protocol). We use the same protocol to prepare both types of GST-RBD 

beads; the only difference is the construct used. The activation assay itself is detailed in 

the Basic Protocol and Alternate Protocol 1. Additional protocols describe how to load 

Rho GTPases with GDP, GTP, or GTPγS (Alternate Protocol 2) and how to perform 

activation assays on nonadherent cells (Alternate Protocol 3). Finally, suggestions on 

how to design activation assays for Rho GTPases other than RhoA, Rac1, and Cdc42 

are given in the Commentary. 

STRATEGIC PLANNING 

The first step consists of making the recombinant GST-RBD fusion proteins. These are 

loaded onto glutathione-coupled beads and stored with the beads at −80◦Cinabuffer containing 10% glycerol. Following this, a small aliquot of the GST-RBD beads is run 

on an SDS-PAGE gel to estimate the amount of fusion protein made and purified. All of 

these steps are described in the Support Protocol. 

The GST-RBD beads then need to be tested to verify that they only interact with the 

active, GTP-bound form of the Rho GTPase of interest. Good controls for Rac1 and 

RhoA activation assays are described in the Basic Protocol. These include treatment 

of serum-starved HeLa cells (70% confluent) with 100 ng/ml epidermal growth factor 

(EGF) for 3 min, which induces robust Rac1 (e.g., see Figure 14.8.2) and RhoA activation. 

A clear increase in active Rac1 or RhoA should be seen when comparing the amount 

of active GTPase in starved and agonist-treated cells. In order to test GST-RBD beads 

before use in Cdc42 activation assays, it is easier to load Cdc42 with GTP than to use 

agonist stimulation because few agonists give marked activation of this small GTPase. 

This method is described in Alternate Protocol 2, and expected results are shown in 

Figure 14.8.4. Since the GST-RBD beads can be stored at −80 ◦ C for long periods of 

time, it is important to test them regularly to ensure they have not lost their specificity 

for active Rho GTPase over time. 

Rac1 ACTIVATION ASSAYS ON HeLa CELLS 

This protocol describes how to prepare the cell lysates to carry out Rac1 and RhoA 

(also see Alternate Protocol 1) activation assays on agonist-treated cells. The conditions 

described in this protocol (see step 2) are used as a quality control or positive control 

for Rac1 or RhoA activation (see Fig. 14.8.2 for an example of a Rac1 activation assay). 

This protocol forms the basis for activation assays that can be carried out on cells 

treated with other agonists or over-expressing Rho GTPase mutants (active, GTP-bound 

or inactive, GDP-bound mutants), inhibitory GTPase activating proteins (GAPs), or 

activating guanine nucleotide exchange factors (GEFs) of interest. This protocol is also 

valid for Cdc42 activation assays although it is unclear which agonist can be used to 

obtain good Cdc42 activation. In this case, the alternative control is to load endogenous 

Cdc42 with GDP, GTP, or GTPγS (see Alternate Protocol 2 and Figure 14.8.4). 

Preparation of cell lysates for Rac1 and RhoA activation assays are identical except 

for the lysis buffers used—lysis buffer B and lysis buffer C (see Alternate Protocol 1), 


espectively. In all cases, it is important to avoid phosphate-based buffers for washing 

or lysing the cells because phosphate forms a precipitate with magnesium (Ren and 

Schwartz, 2000). MgCl2 is present in the lysis buffer to stabilize GTP-bound GTPases 

and to prevent nucleotide exchange. 

In an experimental setting, the level of activation obtained should be quantified by 

normalizing the amount of GST-RBD bound GTPase (i.e., active GTPase) to the amount 

of total GTPase. Care must be taken when quantifying scanned images of X-ray films 

because they have a narrow linear range of exposure (Ren and Schwartz, 2000); it is very 

important to make sure that the films are not overexposed. Activation assays are usually 

performed at least three times so that the mean (± standard deviation) of the ratio of 

active over total GTPase signal can be shown. 

Materials 

HeLa cells: 70% confluent 10-cm plates (two plates per experiment) 


100 µg/ml epidermal growth factor (EGF; Calbiochem); store aliquots at −80◦C DMEM/0.1% (w/v) fatty-acid-free bovine serum albumin (BSA) 

Tris-buffered saline (TBS): 50 mM Tris·Cl (pH 7.6; see APPENDIX 2A)/140 mM 

NaCl, ice cold 

Lysis buffer B, ice cold (see recipe) 

4× SDS-PAGE sample buffer (see recipe) 

GST-PAK-CRIB beads in Tris wash buffer A/10% (v/v) glycerol (Support Protocol) 

Tris wash buffer B (see recipe), ice cold 


SDS-PAGE gel (see UNIT 6.1) 

Refrigerated centrifuge, 4◦C Heating block, 95◦C 0.5- to 20-µl GELoader tips (Eppendorf) 

Additional reagents and equipment for performing SDS-PAGE (UNIT 6.1) and 

immunoblotting (UNIT 6.2) 

1. Wash two 10-cm plates of 70% confluent HeLa cells twice with 10 ml PBS and 

serum starve overnight in DMEM/0.1% (w/v) fatty-acid-free BSA. 

2. Incubate one of the plates of HeLa cells 3 min with 10 ml of 100 ng EGF/ml of 

DMEM/0.1% fatty-acid-free BSA (prepared by diluting the 100 µg/ml EGF in the 

medium). 

The second plate in each experiment is a control. It is serum-starved but not treated with 

agonist. It is incubated overnight in DMEM/0.1% (w/v) fatty-acid-free BSA and remains 

in the cell incubator while treating the other plate with agonist. 

3. On ice, quickly wash the cells twice in 10 ml ice-cold TBS and drain. 

4. Still on ice, extract each plate in 800 µl ice-cold lysis buffer B. 

It is important to keep the samples cold and to work quickly throughout the protocol. 

Over time, the GAP activity in the cell lysates can drastically reduce the amount of active 

GTPase (Ren and Schwartz, 2000). 

5. Centrifuge the cell lysates 10 min at 25,000 × g, 4 ◦ C. Carefully collect the supernatant 

and transfer to a new tube on ice. 

6. Meanwhile, thaw 400 µl GST-PAK-CRIB beads on ice. 

7. Remove 30 µl from each cleared cell lysate for total Rac1 protein control and add 

to 10 µl of4× SDS-PAGE sample buffer. Heat 10 min at 95 ◦ C. 



14.8.3 


Rho GTPase 


14.8.4 

Taking samples from each lysate for each time point is important because these will be 

used to determine the amount of total GTPase, which is used to normalize the amount of 

active, GTP-bound GTPase. 

8. Transfer 650 µl of each cell lysate into a microcentrifuge tube with 200 µl (or the 

equivalent of 30 µg) of GST-PAK-CRIB beads in Tris wash buffer A/10% glycerol. 

9. Rotate 45 min at 4 ◦ C. 

10. Wash the beads four times with 600 µl ice-cold Tris wash buffer B: resuspend 

the beads gently by inversion, centrifuge 20 sec at 500 × g, 4 ◦ C, and remove the 

supernatant. 

11. After the last wash, use the thin GELoader pipet tip to remove all of the remaining 

buffer from the beads. 

This can be done by attaching the GELoader tips to a vacuum aspirator. These tips have 

a smaller diameter than the beads and the beads will not be aspirated. As the buffer is 

removed, the beads should become whiter. 

12. Elute the proteins from the beads by adding 50 µl hot 2× SDS-PAGE sample 

buffer/40 mM DTT and heating 10 min at 95 ◦ C. 

13. Load both the samples for total (obtained in step 7) and active Rac1 (obtained in step 

12) onto an SDS-PAGE gel to resolve the ∼21 kDa GTPase (UNIT 6.1) and process 

for western blotting (UNIT 6.2). 

The amount of total and active Rac1 will vary with the conditions and cell type used, 

so the volume of cell lysate to load in each well should be determined accordingly. As a 

starting point, 20 µl of total and 10 µl of active Rac1 can be loaded for each time point 

(Fig. 14.8.2). 

We have used 15% Tris·Cl gels but have found that 12% Bis-Tris gels run with MES buffer 

give much better resolution and allow for better detection. 

Figure 14.8.2 Rac1 activation assays carried out on HeLa cells. Total Rac1 is shown in the left 

panel and the activation assay in the right panel. After 3 min of EGF treatment (100 ng/ml), Rac1 

is activated as seen by the appearance of Rac1 in the right-hand panel. The upper bands are 

nonspecific signal from the GST-RBD protein. 


MAKING GST-RBD BEADS FOR ACTIVATION ASSAYS 

The CRIB motif of PAK1 interacts with active (i.e., GTP-bound) Rac1 and Cdc42 

(Burbelo et al., 1995) and has thus been used extensively to pull down active Rac1 and 

Cdc42 from cell lysates (Sander et al., 1998; Benard et al., 1999; Benard and Bokoch, 

2002). The WASP CRIB motif has also been used to pull down active Cdc42, but this 

construct does not interact with Rac1 (Haddad et al., 2001). It is still unclear which of the 

CRIB motifs gives a more robust assay for Cdc42 activation. For RhoA, the Rho-binding 

domain of Rhotekin is used (Reid et al., 1996; Ren et al., 1999; Ren and Schwartz, 2000). 

This protocol assumes that the construct of interest is available and already transformed 

into Escherichia coli to make recombinant protein. For activation assays on RhoA, the 

construct used consists of the cDNA sequence for amino acids 7 to 89 of Rhotekin cloned 

into a pGEX2T vector (Amersham; Fig. 14.8.1A). For Rac1 and Cdc42 activation assays, 

we use a construct which consists of the cDNA sequence for amino acids 1 to 253 of 

PAK1 cloned into the same expression vector (Ren and Schwartz, 2000; Fig. 14.8.1B). In 

pGEX vectors the GST-fusion protein is expressed from the tac promoter after induction 

with IPTG. Because these vectors also carry the lacI q gene, they can be used in any E. 

coli host. We have used BL21(DE3)pLysS E. coli, but other hosts such as simple BL21 

E. coli should work as well. 

This protocol describes how to make beads from 4 liters of E. coli culture, which will yield 

enough GST-RBD beads for at least 40 activation assays. The method can theoretically 

be scaled down, but this is not recommended because using smaller volumes may cause 

the solutions to undergo greater temperature fluctuations during the procedure. 

Materials 

LB broth (UNIT 20.2) 

100 mg/ml ampicillin stock solution in water; store small aliquots at −20◦C 34 mg/ml chloramphenicol stock solution in ethanol; store small aliquots at 

−20◦C, protected from light 

E. coli: e. g., BL21(DE3)pLysS (Stratagene) 

GST-RBD construct (obtained from the authors or self-constructed; see Figure 

14.8.1) 

1 M isopropyl-β-D-thiogalactopyranoside (IPTG); store small aliquots at −20◦C Glutathione-Sepharose 4B beads (Amersham) 

Lysis buffer A (see recipe), ice cold 

Tris wash buffer A (see recipe), ice cold 

Tris wash buffer A (see recipe)/10% (v/v) glycerol, ice cold 


12% SDS-PAGE gel (see UNIT 6.1) 

Bovine serum albumin (BSA) 

Coomassie blue staining solution (UNIT 6.1) 

Shaking incubator, 37◦C 2-liter flasks 

Refrigerated centrifuge, 4◦C 30-ml centrifuge tubes, capable of withstanding 17000 × g for 30 min (e. g., Oak 

Ridge), prechilled 

Sonicator 

15- and 50-ml polypropylene tubes (e. g., Falcon), prechilled 

End-over-end rotator, 4◦C Heating block, 95◦C Microcentrifuge tubes, prechilled 

Additional reagents and equipment for performing SDS-PAGE (UNIT 6.1) 

SUPPORT 

PROTOCOL 



14.8.5 


Rho GTPase 


14.8.6 

Grow plasmid-bearing bacteria 

1. Inoculate 5 ml LB broth containing 100 µg/ml ampicillin and 34 µg/ml chloramphenicol 

with a single colony of E. coli containing the GST-RBD construct. 

Instead of keeping glycerol stocks, it is preferred to keep the DNA at −20◦C and to 

transform E. coli when needed. The plates can be kept for a month in the refrigerator. 

Chloramphenicol is required for maintenance of the pLysS plasmid of the 

BL21(DE3)pLysS E. Coli. It is not needed if other hosts are used. 

2. Incubate overnight at 37 ◦ C, with shaking. 

3. Inoculate 400 ml LB broth containing 100 µg/ml ampicillin and 34 µg/ml chloramphenicol 

with the 5-ml overnight culture. 

4. Incubate overnight at 37 ◦ C, with shaking. 

5. Prewarm 4 liters LB broth overnight at 37◦C. LB broth is kept in a 37◦C incubator overnight to save time the next day. 

6. Dilute the overnight culture 1:10 into prewarmed 3.6 liters LB broth containing 100 

µg/ml ampicillin and 34 µg/ml chloramphenicol (typically, 100 ml culture in 900 ml 

broth in a 2-liter flask). 

7. Grow to OD 600 = 0.8. 

This usually takes ∼1 hr, but the OD should be checked after 45 min. 

Induce protein synthesis 

8. Add IPTG to a final concentration of 0.5 mM IPTG using the 1M IPTG stock. 

Incubate 2 hr at 37 ◦ C. 

9. Harvest by centrifuging 25 min at 2500 × g, 4 ◦ C. 

The culture does not have to be precooled. 

10. Carefully remove the supernatant 

11. Keep the bacterial pellets on ice and resuspend them completely in a total of 40 ml 

ice-cold lysis buffer A (i.e., 10 ml lysis buffer A per liter of bacteria culture) 

It is important to keep the solutions cold at all times. Everything should be kept on ice, 

and, if possible, working in a cold room is recommended. 

The bacteria should be completely resuspended before sonication (step 13). This is best 

achieved by first resuspending the pellet into only a small volume of lysis buffer, before 

adding the entire 40 ml. The suspension should be homogenous and devoid of clumps. 

Sonicate bacteria 

12. Transfer the suspension into two prechilled 30-ml centrifuge tubes. 

A volume of ∼20 ml is dispensed equally into each tube. These centrifuge tubes should 

be able to withstand centrifugation at 17,000 × g for 30 min (see step 14). 

13. Sonicate on ice six to eight times for 15 sec each. Cool 2 min on ice between 

sonication bursts to prevent the lysates from overheating. 

It is important that the samples do not overheat and do not foam. The settings for sonication 

will vary from sonicator to sonicator. The samples should be checked regularly. 

Expect the lysate to first become a little viscous as the cells release their DNA—you may 

see a trail from the sonicator probe when you remove it from the lysate. This viscosity 

should disappear with further sonication as the DNA is sheared. It is important that all 

traces of viscous material are removed by sonication, or the lysate will not form a proper 

pellet of debris in the next step. 


14. Centrifuge 30 min at 17,000 × g, 4 ◦ C. 

Prepare beads 

15. In the meantime, equilibrate 0.6 ml (packed bead volume) of glutathione-Sepharose 

4B beads in lysis buffer A by washing them three times in lysis buffer A: add 0.8 

ml lysis buffer A, resuspend the beads by inverting the tube gently, collect the beads 

by brief centrifugation (1 min at 500 × g,4 ◦ C), and remove the supernatant without 

disturbing the bead pellet. 

The beads are sold as a suspension in ethanol. To obtain a reasonably accurate volume 

of packed beads, it is useful to first pipet 0.6 ml of lysis buffer A into a tube (e.g., a 1.5-ml 

microcentrifuge tube) and to draw a line at the level of the meniscus. Remove the buffer 

and add bead suspension until the level of packed beads meets the line on tube. 

Bind GST-RBD to beads 

16. Remove the clarified bacteria lysate supernatant and transfer the ∼40 ml of lysate to 

a prechilled 50-ml polypropylene tube. 

17. Add the equilibrated glutathione-Sepharose 4B beads (prepared in step 15) to the 

lysate. 

To make sure all the beads are transferred, resuspending the beads in a small volume of 

lysate is recommended. The tube that contained the beads can also be washed several 

times with lysate. 


19. Centrifuge 1 min at 500 × g,4 ◦ C. 

Recover bound beads 

20. Carefully discard the supernatant without disturbing the bead pellet and add 12 ml 

ice-cold Tris wash buffer A. 

21. Resuspend the beads by gentle inversions and transfer everything into a prechilled 

15-ml polypropylene tube. Centrifuge 1 min at 500 × g, 4 ◦ C. 

22. Wash the bead pellet another five times by discarding the supernatant, resuspending 

the beads in 12 ml ice-cold Tris wash buffer A by gentle inversions, and centrifuging 

1 min at 500 × g, 4 ◦ C. 

23. Wash once with 12 ml Tris wash buffer A/10% glycerol as in step 22. 

24. Resuspend the beads in 8 ml Tris wash buffer A/10% glycerol. 

25. Remove a 30-µl aliquot of beads in Tris wash buffer A/10% glycerol and add to 

10 µlof4× SDS-PAGE sample buffer. Heat at 95 ◦ C for 10 min and store at −70 ◦ C. 

This sample will be used to check the protein preparation on an SDS-PAGE by Coomassie 

blue staining (see steps 27 to 29). 

26. Divide the beads into small aliquots in prechilled microcentrifuge tubes and store at 

−70 ◦ C. 

200 µl of beads in Tris wash buffer A/10% glycerol are generally used for each activation 

assay; make aliquots of 400 µl and 800 µl for two or four activation assays, respectively. 

Quantify GST-RBD bound to beads 

27. On a 12% SDS-PAGE gel, load 21 µl, 14 µl, and 7 µl of beads in 4× SDS-PAGE 

sample buffer prepared in step 25. 

This is equivalent to loading 15 µl, 10 µl, and 5 µl of beads in Tris wash buffer A/10% 

glycerol. 



14.8.7 


ALTERNATE 

PROTOCOL 1 

Rho GTPase 


14.8.8 

Figure 14.8.3 GST-PAK-CRIB preparation visualized on an SDS-PAGE gel stained with 

Coomassie blue. 45 µl of bead suspension in glycerol buffer was added to 15 µlof4× SDS-PAGE 

sample buffer to give a final volume of 60 µl. Aliquots of 7 µl, 14 µl, and 21 µl (corresponding 

to 5, 10, and 15 µl of bead suspension respectively) were added to each well and compared 

to a dilution range of BSA (10, 5, 2.5, 1, and 0.5 µg per well). In this preparation 5 µl of bead 

suspension is equivalent to ∼2.5 µg ofprotein,sothe200µl of bead suspension used per assay 

is equivalent to ∼100 µg ofprotein(30µg wouldbefine). 

28. Also load different amounts of BSA (e.g., 0.5 µg, 1 µg, 2.5 µg, 5 µg, and 10 µg of 

BSA per well). 

29. Perform electrophoresis, staining with Coomassie blue (see UNIT 6.1). 

30. Estimate the amount of GST-RBD protein bound to the beads by comparing band 

intensities to that of the known amounts of BSA. 

Usually, there is at least ∼2.5 µg of GST-PAK-CRIB for 5 µl of beads in Tris wash buffer 

A/10% glycerol (Fig. 14.8.3). We use 30 to 100 µg of fusion protein per assay. This is 

considerably more than supplied by commercial assay kits, and the assay is consequently 

more sensitive. 

The presence of breakdown products is typical in these preparations, and they do not 

affect the assay (Benard and Bokoch, 2002). 

RhoA ACTIVATION ASSAYS 

The only difference between preparing cell lysates for Rac1/Cdc42 activation assays and 

RhoA activation assays lies in the constituents of the lysis buffer. The lysis buffer used for 

RhoA activation assays (lysis buffer C) contains 0.1% SDS (Ren and Schwartz, 2000). 

This prevents nonspecific interactions in this assay but would affect Rac1 or Cdc42 

binding to GST-PAK-CRIB if used in the Basic Protocol (Benard and Bokoch, 2002). 

As in the Basic Protocol, it is important to avoid phosphate-based buffers for washing 

or lysing the cells because phosphate forms a precipitate with magnesium (Ren and 

Schwartz, 2000). MgCl2 is present in the lysis buffer to stabilize GTP-bound GTPases 

and to prevent nucleotide exchange. The high salt concentration has been shown to help 

minimize GAP activity that might otherwise lead to inactivation of Rho GTPases in the 

lysate (Ren and Schwartz, 2000). 


Additional Materials (also see the Basic Protocol) 

Swiss 3T3 cells: e.g., 70% confluent 10-cm plates (two plates per experiment) 

Lysis buffer C, ice cold 

GST-Rhotekin-RBD beads in wash buffer A/10% (v/v) glycerol (Support Protocol) 

Proceed as in the Basic Protocol with changes in the following steps: 

2. Treat HeLa cells as described in the Basic Protocol, or, alternatively, treat serumstarved 

Swiss 3T3 cells with 10 ml of 1 µg/ml lysophosphatidic acid (LPA) for 1 min 

(Ren et al., 1999) or with 10% (v/v) serum for 2 to 3 min (Ren and Schwartz, 2000). 

4. Still on ice, extract each plate in 800 µl ice-cold lysis buffer C. 

Lysis buffer C contains 0.1% SDS (Ren and Schwartz, 2000). This prevents nonspecific 

interactions, but if used for Rac1 activation assays (see Basic Protocol) would affect Rac1 

or Cdc42 binding to GST-PAK-CRIB (Benard and Bokoch, 2002). 

8. Transfer 650 µl of each cell lysate into a microcentrifuge tube with 200 µl (or the 

equivalent of 30 µg) of GST-Rhotekin-RBD beads in wash buffer A/10% glycerol. 

LOADING Rac1 OR Cdc42 WITH NUCLEOTIDE 

This protocol provides an alternative control to the ones described above and is useful 

when no strong agonist is known (e.g., for Cdc42). The cells are lysed in a buffer devoid 

of magnesium. To load the endogenous GTPases with nucleotide, the cleared lysates are 

spiked with a magnesium-chelating agent (EDTA) and a high concentration of GDP, GTP, 

or GTPγS. Under these conditions, the GTPases will bind the most abundant nucleotide. 

Loading of endogenous GTPases with GTP rather than GTPγS gives a more realistic view 

of the level of activation one can expect in the cell. The exchange is stopped by adding 

magnesium chloride, and GST-RBD beads are added to carry out a pull-down assay. 

An example of a Rac1 and Cdc42 activation assay carried out after nucleotide exchange 

is shown in Figure 14.8.4. An aliquot of the lysate is taken to show basal levels of the 

GTPase of interest before nucleotide exchange. Also recommended is taking an aliquot 

after nucleotide loading (but before adding the beads) to check that the GTPase has not 

been degraded during the incubation at 30 ◦ C. The only difference between the protocols 

for RhoA or Rac1/Cdc42 is the time of incubation at 30 ◦ C, during which nucleotide 

loading takes place. This protocol is theoretically applicable to other Rho GTPases 

but the conditions for nucleotide loading might have to be optimized (e.g., time and 

temperature of incubation). 

Materials 

10-cm plates of cells (two plates for each experiment) 

Tris-buffered saline (TBS): 50 mM Tris·Cl (pH 7.6; see APPENDIX 2A)/140 mM 

NaCl, ice cold 

Lysis buffer D (see recipe), ice cold 


10 mM GTPγS (Calbiochem)/20 mM HEPES, pH 7.4; store aliquots at −20◦C 100 mM GDP (Sigma)/20 mM HEPES, pH 7.4; store aliquots at −20◦C 0.5 M EDTA, pH 8.0 (see APPENDIX 2A) 

1MMgCl2 

GST-RBD beads in Tris wash buffer A/10% glycerol (Support Protocol) 

Tris wash buffer C (see recipe), ice cold 

2× SDS-PAGE sample buffer (see recipe), 95◦C SDS-PAGE gel (see UNIT 6.1) 

ALTERNATE 

PROTOCOL 2 



14.8.9 


Rho GTPase 


14.8.10 

Figure 14.8.4 GDP and GTPγS loading of Rac1 and Cdc42. HeLa cells were lysed and loaded 

with GDP or GTPγS as described in Alternate Protocol 2, and Rac1 or Cdc42 activation assays 

were carried out. The amount of GTPase in lysates to which no nucleotide was added is referred 

to as basal. 

Refrigerated centrifuge, 4◦C Shaking incubator, 30◦C Heating block, 95◦C End-over-end rotator, 4◦C 0.5- to 20-µl GELoader tips (Eppendorf) 

Additional reagents and equipment for determining protein concentration (APPENDIX 

3B or 3H) and performing SDS-PAGE (UNIT 6.1) and immunoblotting (UNIT 6.2) 

1. Wash two 10-cm plates of cells twice with 10 ml cold TBS. 

2. Extract each plate in 600 µl lysis buffer D. 

3. Clear the lysate by centrifuging 10 min at 25,000 × g, 4 ◦ C. 

4. Remove 60 µl of supernatant (to determine total protein; e.g., see APPENDIX 3B or 3H) 

and add 20 µl of4× SDS-PAGE sample buffer. 

5. Transfer 500 µl of each lysate to a new tube. 

6a. To load GTPγS: Add 

6 µlof10mMGTPγS(≥100 µM final concentration) 

12 µlof0.5MEDTApH8.0(≥10 mM final concentration). 

Pipet gently to mix, using a pipet volume of ≥200 µl. 

6b. To load GDP: Add 

6 µl of100mMGDP(≥1 mM final concentration) 

12 µlof0.5MEDTApH8.0(≥10 mM final concentration). 

Pipet gently to mix, using a pipet volume of ≥200 µl. 

7. Incubate 15 min at 30 ◦ C, with gentle shaking (for Cdc42 and Rac1), or 30 min for 

RhoA. 


8. Terminate the exchange by adding 38 µl of 1 M MgCl2 to each tube (≥60 mM final 

concentration). 

9. Add 200 µl (or 30 µg) GST-RBD beads in Tris wash buffer A/10% glycerol to each 

of the tubes. 


11. Centrifuge 20 sec at 500 × g,4 ◦ C, and discard the supernatant. 

12. Wash the beads four times in 600 µl cold Tris wash buffer C: add 600 µl cold Tris 

wash buffer C, invert the tube to gently resuspend the beads, and centrifuge 20 sec 

at 500 × g,4 ◦ C, for each wash. 

13. After the last wash, use the GELoader tips to remove all of the remaining buffer from 

the beads. 

14. Elute with 50 µl of hot (95 ◦ C) 2× SDS-PAGE sample buffer. 

15. Heat 10 min at 95 ◦ C. 

16. Dilute again two-fold in 2× SDS-PAGE sample buffer and load 10 µl per well of an 

SDS-PAGE gel (UNIT 6.1) for immunoblotting (UNIT 6.2). 

The exact amount to load may vary and should be determined accordingly. 

Rac1 ACTIVATION ASSAY ON NONADHERENT CELLS 

Preparing lysates from nonadherent cells to carry out an activation assay consists of 

spiking the cell culture medium with a concentrated solution of agonist and stopping the 

activation by adding an equal volume of 2× lysis buffer. This allows for rapid lysis just 

after agonist treatment. The protocol below is based on that described by Takesono et al. 

(2004). The activation assay for Rac1 is carried out on a lymphocyte cell line treated 

with 100 ng/ml of SDF1α for 0 sec, 30 sec, 2, 5, and 10 min. The results are shown in 

Figure 14.8.5. 

Materials 

Cultures of cells in suspension 

RPMI/0.1% (w/v) fatty-acid-free BSA 

RPMI/0.1% (w/v) fatty-acid-free BSA/20 mM HEPES, pH 7.4 

SDF1α (R&D #350-NS) 

2× lysis buffer (see recipe), ice cold 

GST-PAK-CRIB beads in Tris wash buffer A/10% (v/v) glycerol (Support Protocol) 



SDS-PAGE gel (see UNIT 6.1) 

Refrigerated centrifuge, 8◦C 0.5- to 20-µl GELoader tips (Eppendorf) 

Heating block, 95◦C Additional reagents and equipment for performing SDS-PAGE (UNIT 6.1) and 

immunoblotting (UNIT 6.2) 

Prepare cells for assay 

1. Starve the suspension cells for 4 hr in RPMI/0.1% (w/v) fatty-acid-free BSA. 

It might be possible to starve certain cell lines overnight and this could help reduce the 

basal level of active GTPase. The protocol can be adapted accordingly. 

2. Prewarm the RPMI/ 0.1% (w/v) fatty-acid-free BSA/20 mM HEPES. 

ALTERNATE 

PROTOCOL 3 



14.8.11 


Rho GTPase 


14.8.12 

Figure 14.8.5 Rac1 activation assay carried out on nonadherent cells. Nonadherent 300.19 

cells, a mouse pre-B lymphocyte cell line, were treated with SDF1α for different time points and 

Rac1 activation assays were carried out. The levels of total Rac1 are shown in the top panel 

(exposure of time 30 sec), and the bottom panel shows the levels of active Rac1 (exposure time 

of 1 min 30 sec). Rac1 is activated after 30 sec but the activation is lost by 5 min. 

3. Resuspend the cells at 4.4 × 10 7 cells/ml in RPMI/0.1% (w/v) fatty-acid-free 

BSA/20 mM HEPES. For five time points, prepare 6.6 × 10 7 cells in 1.5 ml. 

4. Transfer 225 µl of cells (10 7 cells) for each of the five time points into microcentrifuge 

tubes. 

5. Prepare at least 120 µl of1µg/ml SDF1α in RPMI/0.1% (w/v) fatty-acid-free 

BSA/20 mM HEPES. 

SDF1α at 1 µg/ml is ten times more concentrated than the final SDF1α concentration 

to which the cells will be exposed. When added to the cells, the concentrated solution of 

SDF1α will be diluted 1:10 by the cell culture medium, and the final concentration of 

SDF1α will be 100 ng/ml. 

6. To the cells corresponding to the 0 sec time point, add 25 µl of RPMI/0.1% (w/v) 

fatty-acid-free BSA/20 mM HEPES. 

Perform assay 

7. To each of the tubes for the other four time points, add 25 µl of the 1 µg/ml SDF1α 

to the 225-µl cell suspension prepared in step 4. 

Each tube will contain 10 7 cells in 250 µl and a final concentration of 100 ng/ml of 

SDF1α. 

8. Stop the reaction by adding 250 µl ice-cold 2× lysis buffer. Keep the lysates on ice. 


Analyze lysates 

9. Clear the lysates by centrifuging 10 min at 25,000 × g, 4 ◦ C. 

10. Meanwhile, thaw 1 ml (or the equivalent of 150 µg) of GST-PAK-CRIB beads in 

Tris wash buffer A/10% glycerol on ice. 

11. Transfer the cleared lysates to new tubes. 

12. For each time point, transfer 30 µl of cleared cell lysate to a new tube and add 10 µl 

of 4× SDS-PAGE sample buffer. Heat 10 min at 95 ◦ C. 

These will be the “total” Rac1 samples. 

13. Add 200 µl of beads in Tris wash buffer A/10% glycerol (or the equivalent of 30 µg) 

to 450 µl of cleared lysate (the rest of the cleared lysate). 

14. Rotate end-over-end 45 min at 4 ◦ C. 

15. Wash the beads four times with 0.5 ml ice-cold 1× lysis buffer (dilute 2× lysis 

buffer): add 0.5 ml cold 1× lysis buffer, invert the tube to gently resuspend the 

beads, and centrifuge 20 sec at 500 × g, 4 ◦ C, for each wash. 

16. After the last wash, use the thin GELoader pipet tip to remove all of the remaining 

buffer from the beads. 

17. Elute the proteins from the beads by adding 50 µl of2× SDS-PAGE sample buffer 

and heat for 10 min at 95 ◦ C. 

18. Load both the samples for total (obtained on step 12) and active Rac1 (obtained 

in step 17) onto an SDS-PAGE gel to resolve the ∼21-kDa GTPase (UNIT 6.1), and 

process for immunoblotting (UNIT 6.2). 




Lysis buffer, 2× (for nonadherent cells) 


300 mM NaCl 

2% (w/v) Triton X-100 

20 mM MgCl2 



Prepare fresh and keep on ice 

Add 0.4 mM PMSF (use 200 mM stock; see recipe) just before use 

For nonadherent cells, the lysis buffer has to be twice as concentrated (2× lysis buffer) 

because it will be diluted in half by the cell culture medium. 

Lysis buffer A 

50 mM Tris·Cl, pH 7.5 (see APPENDIX 2 A) 


150 mM NaCl 

5mMMgCl2 

1mMDTT 





14.8.13 


Rho GTPase 


14.8.14 

Lysis buffer B (for Rac1 & Cdc42 activation) 

50 mM Tris·Cl, pH 7.2 (see APPENDIX 2A) 


500 mM NaCl 

10 mM MgCl2 





Lysis buffer C (for RhoA activation) 



0.1% (w/v) SDS 

500 mM NaCl 

10 mM MgCl2 





LysisbufferD 



500 mM NaCl 





PMSF, 200 mM 

200 mM phenylmethylsulfonyl fluoride in ethanol 

Store protected from light 

SDS-PAGE sample buffer, 4× 

0.5 M Tris·Cl, pH 6.8 (see APPENDIX 2A) 

4% (w/v) SDS 

20% (w/v) glycerol 

40 mM DTT 


SDS-PAGE sample buffer, 2× 


2% (w/v) SDS 

10% (w/v) glycerol 

40 mM DTT 


Note that this sample buffer also contains 40 mM DTT. 

Tris wash buffer A 


0.5% (w/v) Triton X-100 

150 mM NaCl 

continued 


5mMMgCl2 

1mMDTT 


Add 20 µM mM PMSF (use 200 mM stock; see recipe) just before use 

Tris wash buffer B 



150 mM NaCl 

10 mM MgCl2 





Tris wash buffer C 



150 mM NaCl 

30 mM MgCl2 





COMMENTARY 


The Rho proteins belong to the Ras superfamily 

of low-molecular-weight GTPases 

(Wennerberg et al., 2005). They were first 

identified in 1985 (Madaule and Axel, 1985) 

and have now been shown to regulate a wide 

range of cellular processes (Jaffe and Hall, 

2005). Most Rho GTPases cycle between an 

active GTP-bound form and an inactive GDPbound 

form. When bound to GTP, the active 

Rho proteins exert their biological functions 

by interacting with target or effector proteins 

(Bishop and Hall, 2000). In the cell, the GTPases 

are themselves regulated by three different 

classes of proteins. The guanine nucleotide 

exchange factors (GEFs) catalyze the 

exchange of GDP for GTP, thereby activating 

the GTPase (Rossman et al., 2005). The GTPase 

activating proteins (GAPs) stimulate the 

intrinsic GTPase activity of Rho proteins, leading 

to inactivation (Moon and Zheng, 2003). 

Finally, the guanine nucleotide dissociation inhibitors 

(GDIs) affect membrane targeting and 

nucleotide exchange (DerMardirossian and 

Bokoch, 2005; Dovas and Couchman, 2005). 

Activation assays have become fundamental 

to the study of small GTPase function. 

Originally, GTPase activity was measured by 

immunoprecipitating the GTPase of interest 

from lysates of cells that had been metaboli- 

cally labeled with [ 32 P]phosphate and by identifying 

the associated radioactive nucleotide 

using thin layer chromatography (Downward 

et al., 1990). The first nonradioactive activation 

assay was developed for the small GTPase 

Ras. In 1996, Taylor and Shalloway used 

the amino acids 1 to 149 of Raf1 to pull down 

active Ras (Taylor and Shalloway, 1996; 

Taylor et al., 2001). Very soon after, De Rooij 

and Bos (1997) described the same approach 

using a smaller domain of Raf1 (amino acids 

51 to 131). After RhoA was shown to interact 

with Rhotekin in a nucleotide dependent 

fashion (Reid et al., 1996), Martin Schwartz 

and colleagues used the Rho-binding domain 

of Rhotekin to pull down active RhoA 

from cell lysates (Ren et al., 1999; Ren and 

Schwartz, 2000). The CRIB motif of PAK1 

interacts with active (i.e., GTP-bound) Rac1 

and Cdc42 (Burbelo et al., 1995) and has thus 

been used extensively to pull down active 

Rac1 and Cdc42 from cell lysates (Sander 

et al., 1998; Benard et al., 1999; Benard and 

Bokoch, 2002). The WASP CRIB motif has 

also been used to pull down active Cdc42, 

but this construct does not interact with Rac1 

(Haddad et al., 2001). It is still unclear which 

of the CRIB motifs gives a more robust assay 

for Cdc42 activation. 



14.8.15 


Rho GTPase 


14.8.16 

As shown in Figure 14.8.1A, the Rhotekin 

construct used to pull down RhoA usually 

consists of amino acids 7 to 89 (Ren and 

Schwartz, 2000). On the other hand, a range of 

PAK-CRIB constructs have been used successfully 

for Rac1 and Cdc42 activation assays. 

Collard and colleagues used amino acids 56 to 

272 of PAK1B (Sander et al., 1998), whereas 

Bokoch’s group prefers using amino acids 67 

to 150 of PAK1 (Benard and Bokoch, 2002). 

We have used a larger portion of PAK1, namely 

amino acids 1 to 253 (Fig. 14.8.1B). Although 

the CRIB motif is only 16 amino acids long, 

the minimal domain for Rac1/Cdc42 binding 

to PAK consists of residues 75 to 105 

(Thompson et al., 1998). 

Activation assays for small GTPases have 

led to the identification of a wide range of 

extracellular stimuli that result in GTPase 

activation and initiate downstream signaling. 

They are also a good tool for assessing 

basal activation states, especially in tumor 

biology where they can be used to compare 

normal and tumor samples or cell lines. 

GST-RBD pull-down assays are also useful 

for measuring GEF-mediated activation and 

GAP inhibition, but it has been suggested that 

they will not reflect GDI-mediated inhibition 

(Ellerbroek et al., 2003). 

Designing and testing activation assays for 

Rho GTPases other than RhoA, Rac1, and 

Cdc42 

The best characterized Rho GTPases are 

RhoA, Rac1, and Cdc42, but over 20 human 

Rho GTPases have now been identified (Wherlock 

and Mellor, 2002; Wennerberg and Der, 

2004; Boureux et al., 2007). Developing activation 

assays for the other Rho GTPases could 

prove an important tool for studying their function, 

assuming that a specific antibody is available 

for the GTPase of interest. 

Certain Rho GTPases, however, do not 

cycle between an inactive, GDP-bound and 

an active, GTP-bound form. These include 

Rnd1/RhoE, Rnd2, and Rnd3 which are 

thought to be GTPase deficient and constitutively 

GTP-bound (Foster et al., 1996; Nobes 

et al., 1998; Chardin, 2006). Like the Rnd proteins, 

RhoH/TTF, RhoBTB1, and RhoBTB2 

lack the conserved residues corresponding to 

G12 and Q61 (Rac1 numbering) found in 

other Rho GTPases; they are therefore likely 

to be GTPase deficient and not regulated 

by GDP/GTP cycling (Wennerberg and Der, 

2004; Li et al., 2002). Such noncycling Rho 

GTPases are not amenable to activation assays. 

So far, their activity is thought to be regulated 

by their level of expression and/or by phosphorylation 

and could, therefore, be monitored by 

assessing the amount of protein present and/or 

the level of phosphorylation. 

Among the cycling RhoGTPases, RhoB 

and RhoC both bind Rhotekin, and the GST- 

Rhotekin-RBD construct used for RhoA activation 

assays has also been used successfully 

for RhoB and RhoC (Arthur et al., 2002; 

Gampel and Mellor, 2002; Bellovin et al., 

2006; Pan et al., 2006). In this case, it is important 

to ensure that the antibodies used are 

specific and can distinguish between the different 

Rho proteins (Table 14.8.1). The GST- 

PAK-CRIB construct has been used successfully 

to pull down Rac1 and Cdc42-like Rho 

GTPases such as active Rac2 (Benard et al., 

1999) and TC10 (Tong et al., 2007). TCL has 

been shown to bind GST-PAK-CRIB, suggesting 

that this construct could be used to pull 

down active TCL from cell lysates (Vignal 

et al., 2000). Thus, the first step in designing 

an activation assay would be to test whether 

the GST-Rhotekin-RBD, GST-PAK-CRIB, or 

GST-WASP-CRIB constructs can be used (see 

Figure 14.8.1). If this is not the case, one 

needs to identify a protein domain which can 

be made as a GST fusion protein and which 

only interacts with the GTP-bound form of the 

GTPase of interest. 

In order to test whether the GST-fusion protein 

only interacts with the active GTPase, 

pull-downs can be carried out on cells overexpressing 

the inactive, GDP-bound T17N 

mutant or the active, GTP-bound G12V and 

Q61L mutants (Rac1 numbering; Benard et al., 

1999). Alternatively, endogenous GTPases 

can be loaded with nucleotides (Alternate 

Protocol 2). In our experience, not all Rho 

GTPases can be loaded following Alternate 

Protocol 2 because incubation at 30 ◦ C for 15 

or 30 min can lead to degradation of the protein 

of interest. It might be possible to overcome 

this by optimizing the protocol. 

Rho GTPases generally have multiple binding 

partners, with a range of different binding 

parameters. The general sense is that successful 

pull-down assay probes will have high 

affinities for the active Rho GTPase, high 

specificity for the active form, and a low rate 

of dissociation. The last parameter is important 

in preventing loss of the complex during 

the assay procedure. 




When studying agonist-induced activation, 

it is sometimes difficult to obtain low levels of 

active GTPase in the control, untreated cells. 

Not all cell types can be starved and the length 

of serum starvation needed can vary from cell 

type to cell type; this has to be determined empirically. 

The level of active GTPase should be 

as low as possible, and this is usually achieved 

by long periods of serum starvation (e.g., 16 

hr). However, the cells should still look healthy 

after serum starvation. Also, the cells should 

not be too confluent; 60% to 70% confluency 

is usually ideal. Finally, carrying out regular 

quality control activation assays on the GST- 

RBD beads is important. Beads do lose their 

specificity over time and can then interact with 

the inactive form of the GTPase. The only way 

to resolve this is by making new GST-RBD 

beads. 

The major obstacle to activation assays is 

the residual GAP activity found in cell lysates 

(Ren and Schwartz, 2000). The high salt content 

of lysis buffers should help minimize GAP 

activity, but it is also important to work quickly 

and to keep all solutions and samples at low 

temperature (Ren and Schwartz, 2000). If possible, 

it is advisable to carry out the procedure 

in a cold room. Buffers should be made fresh 

and kept on ice. The PMSF is added just before 

use. 

Rho GTPases are not very abundant and 

they can be difficult to detect by immunoblotting. 

Good antibodies are available commercially, 

and these are described in Table 14.8.1. 

The GTPases are only 21 kDa and the SDS- 

PAGE gels used should be chosen accordingly. 

We have used 15% Tris·Cl gels but have found 

that 12% Bis-Tris gels run with MES buffer 

give much better resolution and allow for better 

detection. 

The kinetics of GTPase activation are not 

always predictable. These depend on the cell 

type, the agonist used, and the GTPase studied. 

A typical assay involves a time course of 

0, 30 sec, 1, 3, 5, and 10 min. If no activation 

is seen over this period, it is advisable to use 

a wider range of time points. Activation can 

occur very quickly and be very short lived; alternatively, 

it can occur a very long time after 

treatment with agonist. Indeed, treatment of 

Schwann cells with sphingosine-1-phosphate 

(S1P) leads to maximal RhoA and Rac1 activation 

after only 15 sec (Barber et al., 2004). 

This activation is very rapidly downregulated; 

it is barely noticeable after 1 min and disappears 

completely after 3 min of S1P treatment 

(Barber et al., 2004). On the other hand, treatment 

of HeLa cells with EGF leads to a biphasic 

activation of RhoB, with an early peak in 

activity at 3 min, followed by a second peak 

that appears as late as 60 min after stimulation 

(Gampel and Mellor, 2002). 

In an experimental setting, the level of activation 

obtained should be quantified by normalizing 

the amount of GST-RBD bound GT- 

Pase (i.e., active GTPase) to the amount of 

total GTPase. Care has to be taken when quantifying 

scanned images of X-ray films because 

they have a narrow linear range of exposure 

(Ren and Schwartz, 2000); it is very important 

to make sure that the films are not overexposed. 

Activation assays are usually performed 

at least three times so that the mean 

(± standard deviation) of the ratio of active 

over total GTPase signal can be shown. 


Examples of anticipated results are shown 

in Figures 14.8.2 to 14.8.5. The level of sensitivity 

of the activation assay has not been 

determined. It would vary hugely among cell 

types. Concentrations of agonists used would 

Table 14.8.1 Commercially Available Antibodies for RhoA, RhoB, Rac1, Rac2, and Cdc42 

GTPase Company Catalogue number Species 

RhoAa Santa Cruz RhoA (26C4) sc418 Mouse monoclonal 

RhoBa Santa Cruz RhoB (C5) sc8048 Mouse monoclonal 

Rac1 BD Trans Lab R56220 Mouse monoclonal 

Rac2 Santa Cruz Rac2(C-11) sc96 Rabbit polyclonal 

Cdc42 BD Trans Lab 610928 Mouse monoclonal 

aThe RhoA antibody is specific for RhoA, and similarly, the RhoB antibody only recognizes RhoB and not RhoA or 

RhoC (Gampel and Mellor, 2002). We have not checked the specificity of the other antibodies. 



14.8.17 


Rho GTPase 


14.8.18 

saturate the receptors, based on their known 

affinities. 


The most time consuming task is making 

the GST-RBD beads (Support Protocol). The 

whole procedure is carried out on the same 

day, starting with the inoculation of 4 liters of 

medium with 400 ml overnight culture and finishing 

with the aliquoting and freezing of the 

GST-RBD beads. The beads then need to be 

checked on an SDS-PAGE gel and tested in a 

quality control activation assay using known 

agonists or Rho GTPases loaded with nucleotide. 

Once a batch of beads has been made 

and tested, individual aliquots can be thawed 

on ice just before being added to cell lysates. 

A specific timeline for this process follows: 

After the GST-RBD construct has been made 

or obtained, it needs to be transfected into bacteria 

which are left to grow overnight (day 1). 

A single colony can be picked the next day 

and grown overnight in a 5-ml culture (day 2). 

The 5-ml culture is used to inoculate a 400-ml 

culture, which is grown overnight (day 3). The 

GST-RBD beads (see Support Protocol) can 

then be made in 1 day (day 5) and frozen in 

small aliquots. Checking that the GST-RBD is 

bound to the beads can be done the next day 

(day 6). The beads can be tested as described 

in the Basic Protocol on control cell lysates 

to check that the beads bind the active but not 

the inactive form of the GTPase either on that 

same day (day 6) or the next (day 7). 


Arthur, W.T., Ellerbroek, S.M., Der, C.J., Burridge, 

K., and Wennerberg, K. 2002. XPLN, a guanine 

nucleotide exchange factor for RhoA and RhoB, 

but not RhoC. J. Biol. Chem. 277:42964-42972. 

Barber, S.C., Mellor, H., Gampel, A., and Scolding, 

N.J. 2004. S1P and LPA trigger Schwann cell 

actin changes and migration. Eur. J. Neurosci. 

19:3142-150. 

Bellovin, D.I., Simpson, K.J., Danilov, T., 

Maynard, E., Rimm, D.L., Oettgen, P., and 

Mercurio, A.M. 2006. Reciprocal regulation of 

RhoA and RhoC characterizes the EMT and 

identifies RhoC as a prognostic marker of colon 

carcinoma. Oncogene. 25:6959-6967. 

Benard, V., Bohl, B.P., and Bokoch, G.M. 1999. 

Characterization of rac and cdc42 activation in 

chemoattractant-stimulated human neutrophils 

using a novel assay for active GTPases. J. Biol. 

Chem. 274:13198-13204. 

Benard, V. and Bokoch, G.M. 2002. Assay of 

Cdc42, Rac, and Rho GTPase activation by 

affinity methods. Methods Enzymol. 345:349- 

359. 

Bishop, A.L. and Hall, A. 2000. Rho GTPases and 

their effector proteins. Biochem. J. 348 Pt 2:241- 

255. 

Boureux, A., Vignal, E., Faure, S., and Fort, P. 2007. 

Evolution of the Rho family of Ras-like GTPases 

in eukaryotes. Mol. Biol. Evol. 24:203-216. 

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conserved binding motif defines numerous candidate 

target proteins for both Cdc42 and Rac 

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Chardin, P. 2006. Function and regulation of Rnd 

proteins. Nat. Rev. Mol. Cell Biol. 7:54-62. 

de Rooij, J., and Bos, J.L. 1997. Minimal Rasbinding 

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activation-specific probe for Ras. Oncogene. 

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GTPase activation. Trends Cell Biol. 15:356- 

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Dovas, A., and Couchman, J.R. 2005. RhoGDI: 

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GTPase activities. Biochem. J. 390:1-9. 

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and Cantrell, D.A. 1990. Stimulation of p21ras 

upon T-cell activation. Nature 346:719-723. 

Ellerbroek, S.M., Wennerberg, K., and Burridge, 

K. 2003. Serine phosphorylation negatively regulates 

RhoA in vivo. J. Biol. Chem. 278:19023- 

19031. 

Foster, R., Hu, K.Q., Lu, Y., Nolan, K.M., Thissen, 

J., and Settleman, J. 1996. Identification of a 

novel human Rho protein with unusual properties: 

GTPase deficiency and in vivo farnesylation. 

Mol. Cell Biol. 16:2689-2699. 

Gampel, A. and Mellor, H. 2002. Small interfering 

RNAs as a tool to assign Rho GTPase exchangefactor 

function in vivo. Biochem. J. 366:393- 

398. 

Haddad, E., Zugaza, J.L., Louache, F., Debili, 

N., Crouin, C., Schwarz, K., Fischer, A., 

Vainchenker, W., and Bertoglio, J. 2001. The interaction 

between Cdc42 and WASP is required 

for SDF-1-induced T-lymphocyte chemotaxis. 

Blood 97:33-38. 

Jaffe, A.B. and Hall, A. 2005. Rho GTPases: 

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Biol. 21:247-269. 

Li, X., Bu, X., Lu, B., Avraham, H., Flavell, R.A., 

and Lim, B. 2002. The hematopoiesis-specific 

GTP-binding protein RhoH is GTPase deficient 

and modulates activities of other Rho GT- 

Pases by an inhibitory function. Mol. Cell Biol. 

22:1158-1171. 

Madaule, P. and Axel, R. 1985. A novel ras-related 

gene family. Cell 41:31-40. 

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proteins in cell regulation. Trends Cell 

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Hall, A., and Chardin, P. 1998. A new member 

of the Rho family, Rnd1, promotes disassembly 


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target for Rho bearing homology to a serine/ 

threonine kinase, PKN, and rhophilin in the 

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means go: Turning on Rho GTPases with guanine 

nucleotide-exchange factors. Nat. Rev. Mol. 

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115:239-240. 



14.8.19 


In Vitro GEF and GAP Assays 

Alexander Eberth 1 and Mohammad Reza Ahmadian 1 

1Institut für Biochemie und Molekularbiologie II, Klinikum der Heinrich-Heine-Universität, 

Düsseldorf, Germany 

ABSTRACT 

Small GTPases act as tightly regulated molecular switches governing a large variety 

of critical cellular functions. Their activity is controlled by two different biochemical 

reactions, GDP/GTP exchange and GTP hydrolysis. These very slow reactions require 

catalysis in cells by two kinds of regulatory proteins. While the guanine nucleotide exchange 

factors (GEFs) activate small GTPases by stimulating the slow exchange of bound 

GDP for the cellularly abundant GTP, GTPase-activating proteins (GAPs) accelerate the 

slow intrinsic rate of GTP hydrolysis by several orders of magnitude, leading to inactivation. 

There are a number of methods that can be used to characterize the specificity and 

activity of such regulators, to understand the effect of binding on the protein structure, 

and, ultimately, to obtain insights into their biological functions. This unit describes 

(1) detailed protocols for the expression and the purification of small GTPases and the 

catalytic domains of GEFs and GAPs; (2) preparation of nucleotide-free and fluorescent 

nucleotide-bound small GTPases; and (3) methods for monitoring of the intrinsic 

and GEF-catalyzed nucleotide exchange as well as intrinsic and GAP-stimulated GTP 

hydrolysis. Curr. Protoc. Cell Biol. 43:14.9.1-14.9.25. C○ 2009 by John Wiley & Sons, 

Inc. 

Keywords: fluorescence spectroscopy � guanine nucleotide � mant � tamra 

INTRODUCTION 

A great variety of small GTPases are known, and each of these in turn interacts with a 

variety of regulatory proteins, including guanine nucleotide exchange factors (GEFs) and 

GTPase-activating proteins (GAPs). Analysis of the human genome sequence predicts 

69 GEFs and up to 80 GAPs for Rho family GTPases, which may possibly regulate the 

activity of 19 Rho GTPases. To understand the biological relevance of this molecular 

diversity, it is important to measure the activity and to determine the specificity of 

these regulators in vitro. Only a sparse number of such intermolecular interactions 

have been investigated, primarily using radioactive ligand overlay, filter binding, or 

pull-down assays. These methods are often not sufficient to determine the specificity 

of regulation and quantify the activity of recombinant proteins. However, many of the 

potential interactions defined by these methods require a more detailed analysis of their 

kinetics by appropriate real-time methods. Fluorescent guanine nucleotides are often 

ideally suited to fulfill these criteria, as it is known that they do not grossly disturb the 

biochemical properties of the GTPase and that the fluorescence reporter group is sensitive 

enough to changes in the local environment to produce a sufficiently large fluorescence 

change (Ahmadian et al., 2002; Hemsath and Ahmadian, 2005). Furthermore, it is often 

sensitive to the interaction with partner proteins that happen to bind in its neighborhood. 

This unit describes the role of two different fluorescently labeled guanine nucleotides in 

the biochemical analysis of Rho GTPases (Fig.14.9.1), which can be used to evaluate the 

GEF-catalyzed nucleotide exchange and the GAP-stimulated GTP-hydrolysis activities, 

respectively. Table 14.9.1 describes the protocols presented in this unit. 

Current Protocols in Cell Biology 14.9.1-14.9.25, June 2009 

Published online June 2009 in Wiley Interscience (www.interscience.wiley.com). 

DOI: 10.1002/0471143030.cb1409s43 


UNIT 14.9 





14.9.1 

Supplement 43

BASIC 

PROTOCOL 1 

In Vitro GEF and 

GAP Assays 

14.9.2 

Figure 14.9.1 The chemical structures of the guanosine nucleotide derivatives used in this unit. 

Unlabeled ßuorescent nucleotides contain an OH-group at the position R. 

MEASUREMENT OF INTRINSIC AND SLOW GUANINE NUCLEOTIDE 

EXCHANGE FACTOR (GEF)–CATALYZED NUCLEOTIDE EXCHANGE 

REACTIONS 

To obtain a detailed picture of the molecular switch function of small GTPases and their 

interaction with regulators and effectors, we have established fluorescence-based methods 

for the time-resolved monitoring and quantification of small GTPase functions and 

interactions with their binding partners (Ahmadian et al., 2002; Hemsath and Ahmadian, 

2005). 

Different procedures are available for investigating the guanine nucleotide exchange 

on small GTPases. The dissociation of a protein-bound nucleotide can easily be determined 

in real time by fluorescence spectroscopy using a fluorescent GDP. Usually, 

N-methylanthraniloyl (mant) derivatives of guanosine nucleotides, coupled at the 2 ′ (3 ′ ) 

hydroxyl group of the ribose, are used. 

In principle, each nucleotide-binding protein has a defined intrinsic rate of GDP release, 

which is often too low to be physiologically relevant. Thus, GEFs operate on these small 

GTPases and catalyze the generation of the active GTP-bound state from the inactive 

GDP-bound form. This process is often a result of the GEFs themselves being activated 

or recruited to the vicinity of the corresponding GTPase in response to extracellular 

signaling events. 

Specificity and activity of GEFs can be analyzed qualitatively by comparison of intrinsic 

and GEF-stimulated fluorescence measurements. Usually, this is performed in a fluorescence 

spectrometer, since these reactions are slow (>1000 sec). Here, the bacterially 

expressed recombinant proteins, purified to >90% homogeneity, as well as the chemically 


Table 14.9.1 Protocols for In Vitro GEF and GAP Assays 

Protocol Title 

Basic Protocol 1 Measurement of Intrinsic and Slow Guanine Nucleotide Exchange Factor 

(GEF)–Catalyzed Nucleotide Exchange Reactions 

Support Protocol 1 Preparation of mantGDP-Bound GTPases 

Support Protocol 2 Preparation of Nucleotide-Free Forms of Small GTPases 

Support Protocol 3 Determining Nucleotide Concentration Using HPLC 

Alternate Protocol 1 Measurement of Fast GEF-Catalyzed Nucleotide Exchange Reactions 

Basic Protocol 2 Measurement of GTPase-Activating Protein (GAP)–Stimulated GTP Hydrolysis by HPLC 

Alternate Protocol 2 Measurement of Slow GAP-Stimulated GTP Hydrolysis Using mantGTP 

Alternate Protocol 3 Measurement of Slow GAP-Stimulated GTP Hydrolysis Using tamraGTP 

Alternate Protocol 4 Measurement of Fast GAP-Catalyzed GTP-Hydrolysis Using tamraGTP 

Support Protocol 4 Gene Expression and Bacterial Culture Conditions 

Support Protocol 5 Bacterial Lysis by Sonication 

Support Protocol 6 Bacterial Lysis by a Microfluidizer 

Support Protocol 7 Protein Purification for GST Fusion Proteins 

Support Protocol 8 Determining Protein Concentration Using the Bradford Assay 

Support Protocol 9 Determining Protein Concentration Using the Ehresmann Assay 

Support Protocol 10 Concentrating a Dilute Protein Solution 

Support Protocol 11 Thrombin Proteolytic Cleavage of GST Fusion Proteins 

Support Protocol 12 Gel-Filtration Chromatography 

Support Protocol 13 Freezing and Thawing of Proteins 

synthesized pure nucleotide solution, are prepared in a cuvette. The mant-fluorescence 

signal in a fluorescence spectrometer is recorded using an excitation wavelength of 

366 nm and an emission wavelength of 450 nm, an integration time of at least 2 sec, and 

a recording time for each data point of 20 sec. 

GEF and also GAP assays do not need post-translationally modified GTPases. Thus, 

all proteins and protein domains produced in Escherichia coli can be used. Cleared cell 

lysate is not suitable in this assay for several reasons: (1) protein concentration may not 

be sufficient; (2) the protein of interest may exist in a complex with other proteins and 

may thus not be freely available; (3) the activity of other regulators may falsify the assay. 

Materials 

>5 μM mantGDP-bound GTPase (Support Protocol 1) 

GEF buffer (see recipe), store at 25◦C >50 μM GEF protein including the catalytic domains (recombinant protein, 

expressed and purified as described in Support Protocols 4 to 13) 

10 mM GDP (Pharma Waldhof, http://www.pharmawaldhof.de/), pH 7.5 

0.5 M EDTA, pH 8.0 (APPENDIX 2A) 

Fluorescence cuvettes (Suprasil quartz glass; Hellma, cat. no. 108.002F-QS) 

Fluorescence spectrometer (Perkin-Elmer, Spex Instruments) 

Grafit program (Erithacus Software) or alternative program packages for evaluation 

of the data 

1. Preincubate a solution of 0.1 μM mantGDP-bound GTPase (see Support Protocol 

1) in a fluorescence cuvette in GEF buffer with the GEF protein at a final volume 





14.9.3 


SUPPORT 

PROTOCOL 1 


GAP Assays 

14.9.4 

of 600 μl and at 25 ◦ C for at least 5 min, while measuring the fluorescence signal at 

366 nm excitation and 450 nm emission. 

Always use 366 nm for excitation and 450 nm for emission with mant-labeled nucleotides. 

2. If the fluorescence signal is stable, add 1.2 μl of 10 mM nonfluorescent GDP solution 

(20 μM final GDP concentration) and mix rapidly with a pipet to start the reaction. 

With this setup, the intrinsic dissociation reaction in the absence of GEF is monitored, 

which, depending on the GTPase, lasts between 2 and 72 hr. This slow reaction is 

accelerated in the presence of GEF proteins. To determine the activity and specificity of the 

exchange factors in a fluorimeter, usually concentrations of 0.1 to 2 μM (depending on the 

specific activity of the GEF) should be used (for faster kinetics, see Alternate Protocol 1). 

For slow nucleotide dissociation reactions (>5 hr), the cuvettes should be closed with a 

lid and can additionally be sealed by a thin layer of silicone grease between cuvette and 

lid to prevent evaporation of the sample, which usually results in an artifactual increase 

in fluorescence. 

Alternatively, four parallel measurements can be performed simultaneously using an 

instrument equipped with an automated four-position turret. 

Using cleared cell lysates may disturb the proper fluorescence signal in this assay because 

they are often very opaque solutions and, due to the very low GEF concentrations, useless 

if greatly diluted. 

3. Monitor the exponential decrease in fluorescence over the time course of the reaction 

(2 to 72 hr). 

The decrease in fluorescence is due to the mantGDP release into the aqueous solution. 

4. When no further change in fluorescence can be observed, add 24 μl of 0.5 M EDTA 

solution (for final concentration of 20 mM) and monitor the reaction for an additional 

10 min. 

This will reveal whether the nucleotide dissociation reaction is entirely complete. 

EDTA will deplete the magnesium ion, which is an essential cofactor for nucleotide 

binding, from the nucleotide binding site of the GTPase. This reduces the nucleotide 

affinity by several orders of magnitude and thus leads to a complete dissociation of the 

bound mant-nucleotide. However, the fluorescence baseline will not change if mantGDP 

dissociation is already complete. 

5. Fit the data single exponentially with, e.g., the Grafit program to provide the dissociation 

(off) rates. 

In the case of small GTPases, the dissociation rate is usually ∼10 −3 to 10 −5 sec −1 . 

PREPARATION OF mantGDP-BOUND GTPases 

Loading of nucleotide-free forms of GTPases with fluorescently labeled nucleotides can 

be achieved by simply mixing both components and subsequently performing small-scale 

size-exclusion chromatography with a desalting column. The protocol described below 

is usually necessary for the preparation of GTPases bound to fluorescent GDP analogs. 

For nonhydrolyzable GTP analogs like Gpp(NH)p, the method described in Support 

Protocol 2 is sufficient and, additionally, steps 4 to 5 of Support Protocol 2 could be 

omitted (addition of phosphodiesterase is dispensable). 

Materials 

Standard buffer (see recipe) 

Nucleotide-free GTPase (see Support Protocol 2) 

mantGDP (synthesized as described in Hemsath and Ahmadian, 2005, or 

purchased from Jena Bioscience, http://www.jenabioscience.com/) 


Ponceau S (UNIT 6.2) 

HPLC buffer (see recipe) containing 20% to 25% (v/v) acetonitrile 

NAP-5 column (GE Healthcare) 

Nitrocellulose membrane (UNIT 6.2) 

Additional reagents and equipment for Ponceau S staining of proteins on 

nitrocellulose membrane (UNIT 6.2) and HPLC (Support Protocol 3) 

1. Equilibrate the NAP-5 column with 2 to 3 column volumes of standard buffer. 

2. Mix 0.5 mg of a nucleotide-free GTPase (e.g., 50 μl from a 0.5 mM solution) with 

a 1.5-fold molar excess of mantGDP (e.g., 3.75 μl from a 10 mM stock solution). 

3. Apply complete sample volume to the NAP-5 column and let it sink into the medium. 

4. Add sufficient standard buffer for a 500-μl total buffer/sample volume and let it sink 

into the medium again (e.g., for the example here, add 446.25 μl). 

5. Add 1 ml of standard buffer and collect fractions at 2 drops per fraction. 

Usually the protein elutes in fractions 3 to 5, which corresponds to an elution volume 

between 0.2 and 0.5 ml. 

6. Analyze the fractions for their protein content by dotting 2 μl from each fraction on 

a nitrocellulose membrane and subsequently staining with Ponceau S (UNIT 6.2). 

This is just to determine which fractions contain the protein of interest. 

We do not use any protein standard here since this is not a quantitative analysis. It is 

just qualitative and should answer in which fractions the protein is localized and identify 

which fractions to pool. 

7. Pool protein-containing fractions and determine the mantGDP-bound protein concentration 

by HPLC (Support Protocol 3) using an HPLC buffer containing 20% to 

25% acetonitrile. 

8. Store the protein aliquots at −80 ◦ C. 

It is important to note that the resulting concentration needs to be multiplied by the factor 

0.6 to filter out absorption portions, which trace back to the mant-group absorption at 

254 nm. 

PREPARATION OF NUCLEOTIDE-FREE FORMS OF SMALL GTPases 

Preparation of nucleotide-free GTPase is carried out in two steps according to John et al. 

(1990). In the first step, the bound GDP is degraded by alkaline phosphatase and replaced 

by Gpp(CH2)p (a nonhydrolysable GTP analog, which is resistant to alkaline phosphatase 

but sensitive to phosphodiesterase). In the second step, after GDP is completely degraded, 

snake venom phosphodiesterase is added to the solution of the Gpp(CH2)p-bound GTPase 

to cleave this nucleotide to GMP, G, and Pi. 

Materials 

Alkaline phosphatase, agarose bead-coupled (Sigma-Aldrich) 

Nonhydrolyzable GTP analog Gpp(CH2)p (Sigma-Aldrich) 

GDP-bound GTPase (expressed and purified from E. coli; as described in Support 

Protocols 4 to 13) 

10× exchange buffer (see recipe) 

HPLC buffer (see recipe) containing 7.5% (v/v) acetonitrile 

Snake venom phosphodiesterase (Sigma-Aldrich, cat. no. P3134) 

Liquid nitrogen 

SUPPORT 

PROTOCOL 2 





14.9.5 



GAP Assays 

14.9.6 

Additional reagents and equipment for HPLC (Support Protocol 3 and Basic 

Protocol 2) 

1. Add 0.1 to 1 U of agarose bead–coupled alkaline phosphatase and a 1.5 molar excess 

of Gpp(CH2)p to 1 mg GDP-bound GTPase. 

Use a highly concentrated protein solution of 0.5 to 1 mM in order to obtain highly 

concentrated nucleotide-free GTPases. 

2. Start the reaction by diluting the 10× exchange buffer to 1× in the prepared reaction 

mixture from step 1. Mix exchange buffer and protein/nucleotide-solution rapidly. 

Fast mixing of the exchange buffer should be carried out to prevent local protein precipitation 

due to a high ammonium sulfate concentration. Ammonium sulfate destabilizes the 

nucleotide binding of the GTPase and zinc chloride, and is an essential cofactor of the 

alkaline phosphatase. 

3. Incubate the protein solution at 4 ◦ C for 2 to 16 hr (depending on the GTPase used) 

and analyze the GDP content by HPLC (Support Protocol 3) using an HPLC buffer 

containing 7.5% acetonitrile. 

The GDP peak declines in the course of the degradation progress. It disappears and 

GMP or G peaks appear instead. The amount of Gpp(CH2)p remains unchanged, as it is 

resistant to alkaline phosphatase. 

This protocol can also be used to prepare Gpp(NH)p-bound or mantGp(NH)p-bound 

GTPases as described by Hemsath and Ahmadian (2005). 

4. After GDP is degraded completely, add 0.002 U snake venom phosphodiesterase per 

mg GTPase to cleave Gpp(CH2)p to GMP, G, and Pi. Incubate 2 to 24 hr at 4 ◦ C 

(depending on the GTPase and activity of the phosphodiesterase). 

This is the critical step in the procedure, since here the nucleotide is degraded and 

generation of a low-affinity product can lead to partial denaturation of the protein. In some 

cases, it might be helpful to reduce the amount of exchange buffer used or to do repeated 

cycles of room temperature incubation (e.g., for 20 min) followed by recooling on ice (e.g., 

for 10 to 15 min), which might accelerate the process. The latter method might be employed 

in cases where denaturation of the protein occurs due to a long incubation time (>2 days). 

5. Analyze the degradation process by HPLC (Basic Protocol 2). 

Progress of the reaction can be followed by observing reduction and finally elimination 

of the Gpp(CH2)p peak and an increase in the guanosine peak. 

6. After the degradation of Gpp(CH2)p is complete, centrifuge the solution 2 min at 

1500 × g, 4 ◦ C, to remove bead-coupled alkaline phosphatase and insoluble guanosine. 

Repeat this process two to three times to quantitatively remove all traces of 

alkaline phosphatase-coupled beads, since this enzyme might interfere with subsequent 

biochemical assays performed with the nucleotide-free GTPase. 

Depending on the GTPase, Gpp(CH2)p degradation continues 6 to 18 hr at 4 ◦ C. 

7. Inactivate the phosphodiesterase by snap freezing in liquid nitrogen and quickly 

defrosting at 37 ◦ C, and then repeating the process. Store the protein solution at 

−80 ◦ C. 

Nucleotide-free GTPases are actually in a GMP- and G-bound form that can be stored 

at −80 ◦ C for several months. GMP and G, which are the products of the enzymatic 

degradation of GDP and Gpp(CH2)p, usually exhibit a 6– to 7–orders of magnitude lower 

affinity for the GTPases as shown previously for H-Ras (John et al., 1990). Nonetheless, the 

presence of G and GMP, which can be rapidly exchanged by GDP- and GTP-derivatives, 

provides for stability of the so-called nucleotide-free GTPases. 


DETERMINING NUCLEOTIDE CONCENTRATION USING HPLC 

HPLC allows determination of the concentration of nucleotides (nonlabeled- or mantor 

tamra-labeled nucleotide) using a reversed-phase C18 column (ODS-Hypersil, 5 μM, 

Bischoff Chromatography) and a precolumn (Nucleosil 100 C18, Bischoff Chromatography), 

which separates protein–nucleotide complexes by adsorbing the denatured protein. 

Due to the 1:1 ratio of GTPase and nucleotide complex, this method can be used to 

accurately determine the concentration of a nucleotide-bound protein population. 

Materials 

HPLC buffer (see recipe) 

50 to 100 μM GTPase (recombinant protein, expressed and purified as described in 

Support Protocols 4 to 13) 

Nucleotide standard solutions: e.g., 20 to 400 μM GDP 

Beckman Gold HPLC instrument (Beckman Coulter) 

Reversed-phase C18 HPLC column: Ultrasphere ODS, 5-μM; 250 × 4, 6-mm 

(Beckman Coulter) 

Guard column: Nucleosil 100-5-C18, 5 μM (Bischoff Chromatography; 

http://www.bischoff-chrom.de/) 

20- or 50-μl sample loop (Bischoff Chromatography; http://www. 

bischoff-chrom.de/) 

1. Equilibrate the pump and column of the HPLC system with HPLC buffer until a 

stable baseline at 254 nm is reached. 

For the separation of nonlabeled nucleotides, HPLC buffer (see Reagents and Solutions) 

containing 7.5% acetonitrile is used; 20% to 25% acetonitrile is used for mant- or 

tamra-labeled nucleotides. 

2. Wash the sample loop (20- or 50-μl) with ≥1 ml deionized water. 

3. Prepare ∼30 or 60 μl (depending on the loop size used) of a 50 to 100 μM GTPase 

solution (1 to 2 mg/ml in the case of 21-kDa proteins such as Ras or Rho GTPases) 

in water or standard buffer, and load the entire 20- or 50-μl sample loop with it. 

4. Inject the protein sample prepared in step 3 and monitor eluting nucleotides for 5 to 

10 min (depending on the performance of the reversed-phase column and the flow 

rate; here 1.8 ml/min is used) by absorption at the appropriate wavelength. 

The absorption is detected at 254 nm for nonlabeled as well as mant-labeled nucleotides 

(for the latter, a factor of 0.6 is multiplied by the result, which is based upon the ratio of 

the extinction coefficient of nonlabeled and mant-labeled nucleotides). Molar extinction 

coefficients (at 254 nm) of 13,700 M −1 cm −1 for nonlabeled, 22,000 for mant-labeled, and 

78,000 for tamra-labeled guanine nucleotides (measured at 546 nm) are employed. Due 

to ion–pair formation between the bulky hydrophobic tetrabutylammonium bromide and 

phosphate groups, the elution of the nucleotides is retarded with increasing phosphates. 

Concentrations of tamra-labeled GTPases are not determined by HPLC but spectrophotometrically 

at 546 nm. 

5. Evaluate the nucleotide concentration using a linear absorbance profile of a nucleotide 

standard, e.g., 20 to 400 μM GDP. Calculate the area below a peak in 

the chromatogram using the HPLC integrator and correlate with the absorption of 

the respective probe. Use this value to determine the nucleotide concentration of the 

applied sample by comparison with a linear absorption profile that is generated with 

a nucleotide standard. 

Analogous to the calibration of the Bradford dye solution, the reversed-phase column can 

be calibrated using different GDP nucleotide samples of known concentration. In order to 

calibrate a column, GDP solutions in the range of 20 μM to 400 μM should be prepared, 

SUPPORT 

PROTOCOL 3 





14.9.7 


ALTERNATE 

PROTOCOL 1 


GAP Assays 

14.9.8 

and their concentrations should be verified spectrophotometrically. These samples can be 

then be applied to the column and their respective peak areas can be plotted versus the 

nucleotide concentration. Linear fitting of the data gives a regression line, and the slope 

represents a correlation factor between concentration and peak area. 

MEASUREMENT OF FAST GEF-CATALYZED NUCLEOTIDE EXCHANGE 

REACTIONS 

For fast GEF-catalyzed nucleotide dissociation reactions, the time resolution of a fluorescence 

spectrometer is insufficient for a reliable data analysis. Instead, a stopped-flow 

instrument is routinely used for analysis of rapid kinetics as obtained by quantitative 

GEF-stimulated nucleotide exchange reactions. Here, equal volumes of two different 2× 

samples are automatically injected into a mixing chamber (in a single mixing mode), 

where the fluorescence can be detected directly after the rapid mixing (dead time ∼2 

to 5 msec). Five to eight identical measurements are recorded and averaged in order to 

obtain a higher degree of accuracy. The excitation wavelength for the mant-nucleotides 

is 366 nm and the fluorescence is detected with a cut-off filter mounted in front of a 

photomultiplier (408 nm for mant-nucleotides). 

This is a fast and easy assay to obtain nucleotide exchange activities of GEFs toward 

the respective GTPases. Also, GEFs can be compared in respect to their specificity with 

different GTPases either by simply comparing nucleotide dissociation rates at given 

GEF and GTPase concentrations or by quantitative determination of Michaelis–Menten 

constant (Km) and maximal dissociation rate (kmax). Such kinetic parameters can be 

obtained by using the above condition and increasing concentrations of the GEF proteins 

as described (Guo et al., 2005; Hemsath and Ahmadian, 2005). 


Stopped-flow instrument (Applied Photophysics SX18MV or Hi-Tech SF-61 DX2; 

http://www.photophysics.com) 

NOTE: Because the samples are mixed 1:1, all stock solutions for components of the 

samples should be 2×. 

1. Wash the drive syringes of the stopped-flow instrument several times with 5 to 10 ml 

GEF buffer and adjust the temperature to 25 ◦ C. 

2. Prepare 2× samples in 1× GEF buffer at room temperature (∼25 ◦ C) and a final 

volume of 1000 μl: 

a. One sample contains 0.2 μM of mantGDP-bound GTPase. 

Example: Dilute 2 μl from a 100 μM mantGDP-bound GTPase solution in 998 μl of 

GEF buffer to obtain a 0.2 μM solution of the respective mantGDP-bound protein. 

b. The other sample contains the GEF protein (at a concentrations ranging from 2 

to 1000 μM, depending on the activity and affinity of the GEF for the respective 

GTPase), and 40 μM GDP (200-fold excess above mantGDP). 

A 10-fold excess of the GEF protein usually is a first choice to determine the activity 

of the GEF protein. Therefore, mix 20 μl from a 100 μM GEF solution (20 μM final 

concentration) and 4 μl from a 10 mM GDP solution (40 μM final concentration) in 976 

μl with GEF buffer. 

3. Load each sample into one of the two drive syringes of the stopped-flow instrument. 

Set the excitation wavelength for the mant-nucleotides to 366 nm and detect the 

fluorescence with a cut-off-filter mounted in front of a photomultiplier (408 nm for 

mant-nucleotides). 


Signal-to-noise ratio can be considerably improved if all samples are centrifuged 

(>20 min, 20,000 × g, 4◦C) and if the buffer is filtered and degassed. 

The photomultiplier current should be adjusted manually to a value with a well balanced 

signal-to-noise ratio. More modern instruments are also able to set an optimal value 

automatically. 

4. Start the measurement with the supplied stopped-flow software, which initiates 

pushing the contents of the two syringes containing the samples into the sample cell 

so that both samples join together and rapidly mix at a final volume of about 50 

to 75 μl. Record the fluorescence using excitation at 366 nm and detection with a 

cut-off filter mounted in front of a photomultiplier at 408 nm for mant-nucleotides, 

for a defined time period (depending on the kinetics). 

5. Repeat the mixing and fluorescence recording event up to 10 times until all volume 

in the drive syringe reservoir is consumed. 

Using 1000-μl samples, up to 11 identical measurements can be performed, from which 

the first three are required to equilibrate the sample cell. The fourth measurement can 

be used to reset the photomultiplier current and to determine the timescale for recording 

the complete reaction. Between 400 and 1000 data points should be recorded per measurement. 

Thus, about seven identical measurements can be repeatedly performed and 

averaged to obtain a mean value of all comparable curves. The single exponential data 

obtained corresponds to one averaged experiment at a defined GEF concentration. 

6. Fit all data according to Hemsath and Ahmadian (2005) to obtain the observed rate 

constant (kobs) for the respective concentration of the GEF protein. 

MEASUREMENT OF GTPase-ACTIVATING PROTEIN (GAP)-STIMULATED 

GTP HYDROLYSIS BY HPLC 

All small GTP-binding proteins, with a few exceptions, e.g., Rnd proteins, have a defined 

rate of GTP hydrolysis, which is often too low to be physiologically relevant. Thus, GAPs 

stimulate the very slow intrinsic reaction rate of GTP hydrolysis by several orders of 

magnitude, which is a critical step in signal transduction. 

For the measurement of intrinsic and GAP-stimulated GTP-hydrolysis reaction of small 

GTPases, several assays have been developed so far. There are filter-binding and HPLCbased 

assays as well as spectrophotometric and spectrofluorometric methods available. 

A generally useful and accurate method is HPLC, by which concentrations of GDP and 

GTP can be determined to describe the reaction progress as described for Ras (Ahmadian 

et al., 1999) and Rho-proteins (Hemsath and Ahmadian, 2005). 

HPLC needs large amounts of proteins and is not suitable for quantitative analysis of 

the GAP activity. In another approach, which is less material- and time-consuming, 

the GTPase reaction rates can be conveniently measured in real time using tryptophan 

fluorescence with an excitation wavelength of 295 nm and an emission wavelength of 

350 nm. For example, the Ras(Y32W) mutant provides a large increase in fluorescence 

signal upon hydrolysis of GTP to GDP and inorganic phosphate, which has been used to 

study the mechanism of the intrinsic GTPase reaction (Ahmadian et al., 1999). 

Stopped-flow experiments using mantGTP and mantGpp(NH)p (see Alternate Protocol 

2) have provided key insights into the mechanism of the GAP-stimulated GTPase reaction 

of Ras in several studies (Ahmadian et al., 1997a,b, 2003). However, it should be noted 

that mantGTP does not work for most GTPase/GAP systems. A case in point is the 

interaction between Rho and RhoGAP, which, although similar in the type of enzymatic 

catalysis to the Ras–RasGAP interaction (Scheffzek and Ahmadian, 2005), does not 

show any fluorescence change upon interaction or hydrolysis. 

BASIC 

PROTOCOL 2 





14.9.9 



GAP Assays 

14.9.10 

Most recently, we have demonstrated that tetramethylrhodamine (tamra; see Alternate 

Protocol 4) is a powerful fluorescence reporter group to study the GTP-hydrolysis in the 

presence and in the absence of GAPs (Eberth et al., 2005). The intrinsic GTP-hydrolysis 

reaction of Ras and Rho GTPases can be detected in a fluorescence spectrometer or a 

stopped-flow instrument, showing a significant decrease in the fluorescence signal. 

The HPLC method is an accurate way of determining the intrinsic hydrolysis rate of 

a GTPase and is also useful for determining a GAP protein’s specific activity. The 

advantage is the lack of any artificial reporter group or necessity to use a GTPase mutant 

in this assay. Here wild-type proteins and unlabeled nucleotides can be used. 

Materials 

500 μM nucleotide-free GTPase (Support Protocol 2) 

GAP buffer (see recipe) 

10 mM GTP (Pharma Waldhof; http://www.pharmawaldhof.de/), pH 7.5 

>50 μM GAP protein including the catalytic domains (recombinant protein, 

expressed and purified as described in Support Protocols 4 to 13) 


HPLC buffer (see recipe) containing 7.5% (v/v) acetonitrile 

Thermomixer/thermoblock 

Beckman Gold HPLC instrument (Beckman Coulter) 

Reversed-phase C18 HPLC column: Ultrasphere ODS, 5-μM; 250 × 4.6–mm 

(Beckman Coulter) 

Guard column: Nucleosil 100-5-C18, 5-μM (Bischoff Chromatography) 

1. Dilute 40 μl of 500 μM nucleotide-free GTPase with 60 μl GAP buffer and the 

GAP protein and preincubate it at 25 ◦ C in a thermomixer/thermoblock (the final 

concentration of the nucleotide-free GTPase should be 80 μM in a volume of 250 μl 

after addition of the GTP). 

The GAP protein can be used in a ratio of 1:100 to 1:1000 with respect to the GTPase 

concentration (0.8 to 0.008 μM in this example) and should be added in step 1 to the 

nucleotide-free GTPase before starting the reaction by addition of GTP. 

2. Dilute 10 μl of 10 mM GTP stock solution with 990 μl GAP buffer (1:100 dilution) 

to obtain a 100 μM GTP solution, and preincubate this solution at 25 ◦ Cina 

thermomixer/thermoblock. 

3. Add 150 μl of the 100 μM GTP solution (60 μM final concentration) and mix it with 

the 80 μM nucleotide-free GTPase solution by pipetting up and down. 

The total volume (in this case calculated for 8 time points) should be 250 μl, but depending 

on the number of data points to be taken, can be increased or even decreased. 

The timescale for the intrinsic GTP hydrolysis reaction varies from 0.5 to 6 hr, which 

depends on the GTPase variant used. Alternatively, a protein comprising a catalytic active 

GAP domain can be added before starting the reaction, which allows the determination 

of the activity and specificity of the GAP protein. 

Note that in the presence of the GAP, the reactions are much faster and the time scale (e.g., 

1 to 5 min instead of 30 to 60 min for RhoGTPases) is much shorter. Using the conditions 

described here, the reaction is usually accelerated by a factor of 5 to 20 (depending on 

the dilution ratio of GAP with respect to GTPase and the activity of the GAP protein). 

In order to obtain reliable results, parameters including GAP concentration and the time 

points need to be optimized for the respective GTPase/GAP system. 

In the case of slow GTP-hydrolyzing GTPases (e.g., Ras or Rap) or GTP-hydrolysis, 

deficient Rho or Ras proteins GTP-bound proteins can be prepared and stored at −80 ◦ C 

for several weeks. 


Depending on the sensitivity of the HPLC, the concentrations of GTPase and GTP can be 

reduced to 25 μM and 20 μM, respectively. 

4. At defined time points, collect and immediately snap-freeze 30-μl samples in liquid 

nitrogen. 

Proper time points and intervals are important for an appropriate evaluation of the data 

to determine the rate constant of the hydrolysis reaction. These need to be adjusted to the 

activity of the respective GTPases. In principle, 7 to 8 time points are enough to cover a 

complete reaction. The intervals increase in the course of the reaction. For example 0, 1, 

2, 5, 10, 15, 20, and 30 min have been used to determine the GTP hydrolysis reaction of 

the Rac proteins (Haeusler et al., 2003) and 0, 5, 10, 15, 20, 30, 40, 80, 150, and 200 min 

to determine that of H-Ras wild-type (Ahmadian et al., 1999). 

5. Defrost each sample of the defined incubation interval at 95 ◦ C (using a thermoblock 

for about 10 sec) such that they are freshly thawed just before injecting on a reversedphase 

analytical HPLC system. 

The HPLC system should be equipped with a guard column in between the injector 

valve and the analytical C18 reversed phase column. The buffer condition used for the 

separation of non-fluorescently-labeled nucleotides is HPLC buffer (see Reagents and 

Solutions) containing 7.5% acetonitrile. The retention time of guanine nucleotides on 

such an HPLC system and under the described isocratic conditions correlates with the 

number of phosphate groups of the nucleotide (retention order: GMP, GDP, and then 

GTP). 

6. Determine the amount of each nucleotide immediately, using the HPLC integrator, 

from the area integration of the GTP- and GDP-peaks, respectively. 

These values are used to calculate the relative content of the GTP-bound GTPase species 

from the ratio (GTP)/(GTP) + (GDP). 

GTP content (y-axis) can be plotted versus the incubation time of the respective sample 

(x-axis) and fitted with a single exponential. Intrinsic GTP-hydrolysis rates of small 

GTPases usually are in the region of 0.0005 to 0.5 min−1 . 

MEASUREMENT OF SLOW GAP-STIMULATED GTP HYDROLYSIS USING 

mantGTP 

Since there is almost no difference in the fluorescence signal between mantGTP and 

mantGDP, mantGTP-hydrolysis by the GTPase cannot be monitored directly. However, 

association of GAPs with the mantGTP-bound GTPase provides a tool to measure 

GAP-stimulated GTP-hydrolysis reaction, as shown for the Ras and RasGAP proteins 

(Ahmadian et al., 1997a). The GAP-stimulated GTP hydrolysis can be recorded with 

a stopped-flow instrument as a consequence of rapid association and dissociation of 

the GAP protein. Since this assay accounts only for the Ras/RasGAP system, here, the 

specific proteins Ras and the catalytic domain of p120RasGAP are mentioned as an 

example. 

Additional Materials (also see Basic Protocol 2 and Alternate Protocol 1) 

10 mM mantGTP (synthesized as described in Hemsath and Ahmadian, 2005 or 

purchased from Jena Biosciences) bound to Ras, pH 7.5 

20 μM catalytic domains of RasGAP protein (e.g., p120RasGAP )oranyotherGAP 

protein specific for Ras (expressed and purified as in Support Protocols 4 to 13) 

Stopped-flow instrument (Applied Photophysics SX18MV or Hi-Tech SF-61 DX2) 



1. Wash the drive syringes of the stopped-flow instrument several times with 5 to 10 ml 

GAP buffer and adjust the temperature to 25 ◦ C. 

ALTERNATE 

PROTOCOL 2 





14.9.11 


ALTERNATE 

PROTOCOL 3 


GAP Assays 

14.9.12 

2. Charge one drive syringe with 2 μM mantGTP-bound Ras and fill the other syringe 

with 20 μM catalytic domains of the RasGAP protein (e. g., p120 RasGAP ) in GAP 

buffer at 25 ◦ C. 

3. Set the excitation wavelength to 366 nm (as for all mant-nucleotides) and restrict the 

emission light to >408 nm by a corresponding cut-off-filter. 

The same rules concerning volumes to prepare and photmultiplier settings described in 

Alternate Protocol 1 also apply to this assay. 

4. After the initial injections to prepare the system for reliable data recording, carry out 

individual measurements, which usually show a rapid decrease of fluorescence. 

Change in fluorescence is, in this case, due to the GAP dissociation from the GDPbound 

Ras after GTP hydrolysis. This implies a preceding fluorescence increase upon 

GAP association with the GTP-bound Ras, which cannot be resolved temporally by the 

stopped-flow machine due to the large rate constant of the association process. Depending 

on the rate constant of the association it is possible to observe (by a stopped-flow system 

with a short dead time) both association and GTP hydrolysis in one experiment, following 

subsequent dissociation of the Ras-RasGAP complex, which allows mechanistic studies 

(Ahmadian et al., 1997a,b). 

These measurements are in principle performed using the same procedure and conditions 

described in Alternate Protocol 1. 

MEASUREMENT OF SLOW GAP-STIMULATED GTP HYDROLYSIS USING 

tamraGTP 

In contrast to other fluorescent nucleotide derivatives, including the mant-nucleotides, 

tamraGTP (a ribose hydroxyl-substituted tetramethylrhodamine derivative of GTP) enables 

us to measure the intrinsic and GAP-stimulated GTPase reactions of Rho and 

Ras proteins using fluorescence spectroscopy (Eberth et al., 2005). Besides much lower 

consumption of proteins and nucleotides as compared to the HPLC-based assay, the 

tamraGTP hydrolysis assay allows monitoring the real-time kinetics of the hydrolysis 

reaction of the Ras and Rho families. Intrinsic and GAP-stimulated hydrolysis reactions 

are recorded by a fluorescence spectrometer at an excitation wavelength of 546 nm and an 

emission wavelength of 583 nm, with an integration time of at least 2 sec and a recording 

time for each data point of 20 sec. 


2 mM tamraGTP (synthesized as described in Eberth et al., 2005), pH 7.5 

50 μM nucleotide-free GTPase (Support Protocol 2) in GAP buffer (see recipe for 

buffer) 

Fluorescence cuvettes, Suprasil quartz glass; Hellma, cat. no. 108.002F-QS 

Fluorescence spectrometer (Perkin Elmer, Spex Instruments) 

Grafit program (Erithacus Software) or alternative program packages for evaluation 

of the data 

1. Preincubate a solution of 0.1 μM tamraGTP (prepared from 2 mM stock solution) 

in a fluorescence cuvette in GAP buffer (stored at 25 ◦ C) with the GAP protein, at a 

final volume of 600 μlat25 ◦ C for at least 5 min. 

The GAP protein would be added in step 1 to the tamraGTP before the reaction is started 

by addition of nucleotide-free GTPase in step 2. 

2. Add 1.8 μl of50μM stock solution of nucleotide-free GTPase (0.15 μM final 

concentration), to observe complex formation with the nucleotide through a strong 

increase in fluorescence. 

Alternatively, four parallel measurements can be performed simultaneously using an 

instrument equipped with an automated four-position turret. 


3. After this initial phase of nucleotide association, monitor the significant fluorescence 

decay as a result of GTP hydrolysis, which lasts between 0.5 and 6 hr, depending on 

the GTPase variant used (Eberth et al., 2005). 

Such an intrinsic reaction is significantly faster in the presence of a catalytic amount of a 

GAP protein (0.001 μMto1μM, depending on the specific activity of the GAP protein). 

TamraGTP fluorescence cannot be used for Rab, Ran, alpha-subunits of the heterotrimeric 

G-proteins, and elongation factors (Eberth et al., 2005). 

4. Continue the measurement until no further decrease in fluorescence can be observed. 

5. Evaluate data obtained by single-exponential fitting with a scientific software, e.g., 

the Grafit program. 

MEASUREMENT OF FAST GAP-CATALYZED GTP HYDROLYSIS WITH 

tamraGTP 

The following protocol measures GAP-stimulated tamraGTP hydrolysis by a stoppedflow 

instrument as described in Alternate Protocol 1, except that an excitation wavelength 

of 546 nm is employed and a cut-off-filter of 570 nm in front of a photomultiplier is used 

to detect the fluorescence. 

Additional Materials (also see Basic Protocol 2 and Alternate Protocol 1) 

tamraGTP (synthesized as described in Eberth et al., 2005; 2 mM in deionized 

H2O, pH 7.5) 

Stopped-flow instrument (Applied Photophysics SX18MV or Hi-Tech SF-61 

DX2) 



1. Wash the drive syringes of the stopped-flow instrument several times with 

5 to 10 ml GAP buffer and adjust the temperature to 25 ◦ C. 

2. Prepare 2× samples in GAP buffer at room temperature (∼25 ◦ C) and a final volume 

of 1000 μl: 

a. One sample contains 0.3 μM nucleotide-free GTPase and 0.2 μM tamraGTP. 

b. The other sample contains the GAP protein (at a concentration ranging from 0.2 to 

200 μM, depending on the activity and affinity of the GAP for the respective 

GTPase). 

Example: (1) prepare the GAP solution by diluting the concentrated GAP protein to a 

defined concentration between 0.2 to 200 μM in GAP buffer. (2) Dilute 6 μl froma 

50 μM solution of nucleotide-free GTPase (0.3 μM final concentration) in 984 μl GAP 

buffer and add 10 μl froma20μM tamraGTP solution (0.2 μM final concentration) 

just prior to loading the drive syringes of the stopped-flow instrument with your sample. 

The latter sample is prepared immediately before starting the measurement in order to 

avoid intrinsic tamraGTP-hydrolysis, particularly in the case of rapidly GTP-hydrolyzing 

proteins such as Rho GTPases. 

In the case of slow GTP-hydrolyzing GTPases (e.g., Ras or Rap) or GTP-hydrolysisdeficient 

Rho or Ras proteins, tamraGTP-bound proteins can be prepared and stored at 

−80◦C for several weeks. 

Using 1000-μl samples, up to 11 identical measurements can be sequentially performed, 

from which the first three are required equilibrate the sample cell. The fourth measurement 

can be used to reset of the photomultiplier current and to determine the timescale for 

recording the complete reaction. 

ALTERNATE 

PROTOCOL 4 





14.9.13 


SUPPORT 

PROTOCOL 4 


GAP Assays 

14.9.14 

The photomultiplier current should be adjusted manually to a value with a well-balanced 

signal-to-noise ratio. Alternatively, modern instruments are also able to set an optimal 

value automatically. 

The single-exponential data obtained corresponds to one experiment at a defined GAP 

concentration. 

To obtain kcat and Kd values, increase GAP concentrations as described (Eberth et al., 

2005). 

3. Continue as described in steps 3 to 6 of Alternate Protocol 1, except use 546 nm as 

excitation wavelength and a 570 nm cutoff filter mounted in front of the photomultiplier. 

GENE EXPRESSION AND BACTERIAL CULTURE CONDITIONS 

For the optimization of synthesis of a protein of interest in E. coli, various culture conditions 

should be examined including concentration of isopropyl-D-thiogalactopyranoside 

(IPTG; the inducer of lac-promoter-controlled gene expression), optical density of culture 

at IPTG induction, and, particularly, the temperature and culturing time. These 

culture-condition tests should be performed in small-scale (20 to 50 ml) pilot studies 

prior to scaling up to large-scale cultures for preparative protein expression. To improve 

the maximal recovery, we alternatively used, besides E. coli strain BL21 (DE3), strains 

containing additional plasmids such as BL21 pLys S (to improve bacterial lysis and 

for the expression of toxic proteins), and BL21 Codon Plus RIL or Rosetta (DE3) (to 

improve the codon usage). 

All recombinant proteins described in this unit, including GTPases, GEFs, GAPs, and 

their catalytically active fragments are expressed in E. coli and purified using the methods 

described in this and the following support protocols. 

Materials 

Escherichia coli strain: BL21(DE3), BL21(DE3) Codon plus RIL, BL21(DE3) 

pLysS, or Rosetta (DE3) (Novagen) containing prokaryotic expression plasmid 

carrying gene for protein of interest 

Terrific broth medium (TB medium; see recipe) 

Appropriate selection antibiotics 

Isopropyl-β-D-thiogalactopyranoside (IPTG; Gerbu Biochemicals, 

http://www.gerbu.de) 

Wash buffer (see recipe) 

150- to 1000-ml and 5-liter sterilized Erlenmeyer flasks 

Horizontal environmental shaker incubator (Infors HT, http://www.infors-ht.com/) 

1000-ml and 30- to 250-ml centrifuge bottles 

Centrifuge: Avanti J-20 XP (Beckman Coulter) or equivalent 

6-liter rotor: JLA-8.1000 (Beckman Coulter) or equivalent 

50-ml plastic tubes 

1. Grow 30 to 250-ml precultures of the desired E. coli strain in TB medium in a 150to 

1000-ml flask overnight at 37 ◦ C. 

Remember to add the required antibiotics to the TB medium to maintain transformed 

plasmids. If using BL21 (DE3) Codon plus RIL, BL21 (DE3) pLysS, or Rosetta (DE3) 

strains, chloramphenicol (25 mg/liter) needs to be added. 

2. Fill each 5-liter Erlenmeyer flask to be used with 2.5 liter TB medium. Inoculate 

each flask with 25 ml of an overnight preculture. 

Cultivations usually are carried out in 2.5- to 20-liter scale, depending on the expression 

level and yield of the particular protein. 


3. Place the inoculated culture flasks in a horizontal environmental shaker and let them 

grow at 37 ◦ C and 160 rpm. 

4. When the logarithmic growth phase is reached (OD600 = 0.4 to 0.8), lower the 

temperature to the previously determined optimal expression condition (usually 18 ◦ 

to 30 ◦ C), and add IPTG (usually 0.05 mM to 0.5 mM; depending on pilot tests of 

expression conditions) to induce gene expression. 

The small GTPases as well as GEF and GAP proteins including their catalytic domains 

are usually expressed at an optical density of OD600 = 0.6 to 0.8, with 0.1 mM IPTG, and 

at 18 ◦ to 25 ◦ C overnight. 

5. Perform the incubation either overnight, or for only 2 to 4 hr in difficult cases, 

depending on the results from the optimization of expression tests. 

6. Transfer the cells to 1000-ml centrifuge bottles and harvest the cells by centrifugation 

for 15 min at 6000 × g,4 ◦ C, using a 6-liter rotor if available. Repeat this step several 

times if the culture volume exceeds the capacity of the available rotor. 

7. Wash the bacterial pellet in each rotor bottle with 20 ml wash buffer. Combine 

the resuspended cell pellets into a smaller (30- to 250-ml) rotor bottle (tared) and 

centrifuge again for 20 min at 6000 × g, 4 ◦ C. 

This step is carried out in order to remove residual medium. 

8. Discard the supernatant and weigh the bacterial pellet–containing bottle. Determine 

the weight of the bacterial pellet and resuspend it in wash buffer (3 ml/g bacterial 

pellet) or any other proper buffer that is suitable for solubilizing and stabilizing the 

desired protein. Prepare aliquots in 50-ml plastic tubes. 

9. Store aliquots at −20 ◦ C. 

In addition to cryopreserving the sample, freezing will also help to improve the efficiency 

of bacterial lysis. 

BACTERIAL LYSIS BY SONICATION 

An efficient bacterial lysis is an important prerequisite for the complete recovery of 

expressed recombinant protein. Cell walls of bacteria which have produced proteins of 

interest must be disrupted in order to allow access to intracellular components. Different 

methods have been developed to achieve this goal; they vary considerably in the severity of 

the disruption process, reagents needed, and the equipment available. Besides enzymatic 

methods using lysozyme treatment, which is suitable for analytical scale and not always 

reproducible, there are several mechanical methods—including glass bead, cell bomb, 

French press, sonication, and microfluidizer—to gently disrupt bacterial cell walls. We 

commonly use the latter two methods, both of which are efficient and fairly quick. This 

protocol describes sonication, which permits cell disruption in smaller samples (≥200 μl 

and ≤200 ml). 

Materials 

70% ethanol 

Bacterial sample (Support Protocol 4) 

Pefabloc (ICN Biochemicals) 

Lysozyme (Sigma-Aldrich) 

DNase I (Sigma-Aldrich) 

Sonicator: Branson Sonifier S-450A and 3- to 19-mm titanium probe 

NOTE: The protease inhibitor Pefabloc (0.02% w/v) and lysozyme (2 μg/ml suspension), 

as well as DNase I (10 μg/ml suspension) are added to the bacterial suspension before 

lysis. 

SUPPORT 

PROTOCOL 5 





14.9.15 


SUPPORT 

PROTOCOL 6 


GAP Assays 

14.9.16 

1. Equip the cell sonicator with a titanium probe of 3- to 19-mm diameter (depending 

on the culture volume to be disrupted) and clean it before use with 70% ethanol. 

Use a 3-mm sonifier probe for volumes of 2 to 50 ml and a 19-mm probe for volumes of 

50 to 500 ml. 

2. Transfer the thawed bacterial suspension from all aliquots to a beaker of suitable 

size and place it on ice. 

3. Place the sonicator probe about 0.5 to 1 cm beneath the surface into the suspension. 

4. Start the sonication procedure by increasing the output control (add 5 to 10 W 

each time) at 10-sec intervals starting with 30 W and ending with ∼95 W. Repeat 

this procedure eight to twelve times, and always wait 30 sec in between to prevent 

overheating of the sample. Keep the beaker with the bacterial solution on ice during 

the entire procedure. 

Optionally, the wave duty cycle function of the ultrasonic instrument can be used to reduce 

heat production and free radical formation. 

A color change from very milky to slightly more translucent should be observed and can 

act as an indicator for cell disruption. 

BACTERIAL LYSIS USING A MICROFLUIDIZER 

The microfluidizer is an instrument that uses high pressure to squeeze the bacterial 

solution through a flux cell containing a narrow channel, thereby generating high shear 

forces that pull the cells apart. The system permits controlled cell breakage and does 

not require addition of detergent or higher ionic strength. Since heat is generated during 

this process, the flux cell needs to be cooled. The microfluidizer system provides a 

convenient and efficient method for cell lysis of larger cell suspensions (≥20 ml to several 

liters). 

Materials 

Bacterial sample (Support Protocol 4) 

Buffer to be used for protein purification 

Pefabloc (ICN Biochemicals) 

Lysozyme (Sigma-Aldrich) 

DNase I (Sigma-Aldrich) 

100% 2-propanol 

Microfluidizer (Microfluidics Corp., http://www.microfluidicscorp.com) 

NOTE: The protease inhibitor Pefabloc (0.02% w/v) and lysozyme (2 μg/ml suspension) 

as well as DNase I (10 μg/ml suspension) are added to the bacterial suspension before 

lysis. 

1. Wash the instrument extensively with water. For a final wash step, use the standard 

buffer for the protein purification. 

This will remove all traces of alcohol in which the instrument is usually stored to prevent 

microbial growth. 

2. Pour the thawed bacterial suspension into the instrument’s reservoir and turn on the 

instrument 

Direct the flow of the instrument’s outlet toward the wall of a beaker to prevent foam 

formation. 

3. Prevent intake of air on the inlet, as this will also produce foam and lead to protein 

denaturation. For this, turn off the instrument before air enters the instrument’s inlet. 


Wash with a small volume of standard buffer and switch the instrument on again. 

Stop again before air enters the inlet of the instrument. 

By repeating this step two to three times, nearly all of the bacterial suspension will be 

processed. 

4. If necessary, flush the bacterial solution two to three times through the instrument, 

until a color change from milky to slightly more translucent is observed. 

5. Wash the instrument extensively with water and finally with 2-propanol, and store it 

in this solution. 

PROTEIN PURIFICATION FOR GST FUSION PROTEINS 

The use of proteins with high purity (>95%) is a mandatory prerequisite for investigation 

of protein structure-functional relationships. The use of recombinant protein expression 

systems and the development of a variety of fusion tags has facilitated the preparation of 

high-purity proteins dramatically. Nevertheless, choosing the right purification strategy 

is still a matter of trial and error and has to be discovered for each individual protein. 

The GST-fusion affinity purification system is a well established technique and works 

successfully for purification of GTPases and their regulatory proteins. 

Materials 

Bacterial lysate with an overexpressed GST-fusion protein 

Glutathione-Sepharose 4B FF (GE Healthcare) 


Standard buffer (see recipe) containing 500 mM KCl and 1 mM ATP 

Standard buffer containing 20 mM glutathione (adjusted to pH 7.5 with NaOH 

again after addition of glutathione) 

0.01% (w/v) sodium azide or 20% (v/v) ethanol 

Centrifuge: Avanti J-30I (Beckman Coulter) or equivalent 

Rotor: JA-30.50 or JA-17 (Beckman Coulter) or equivalent 

Äkta Sytem, e.g. Äkta Prime (GE Healthcare) 

XK 26/20 chromatography column chassis (GE Healthcare) 

Additional reagents and equipment for SDS-PAGE (UNIT 6.1) and Coomassie 

staining (UNIT 6.6) 

Centrifuge lysate 

1. Separate insoluble constituents from the bacterial lysate by centrifuging 40 min at 

35,000 to 100,000 × g,4 ◦ C. 

If possible, centrifuge at 100,000 × g to remove as much insoluble cell debris as possible. 

If such a high-speed rotor/centrifuge is not available, a minimal force of 35,000 × g might 

also be sufficient. 

Prepare glutathione-Sepharose column 

2. Equilibrate an XK 26/20 column packed with Glutathione-Sepharose (≥25-ml bed 

volume) with ∼3 to 4 column volumes of standard buffer until a stable baseline is 

reached. Monitor absorption at 280 nm using the Äkta System. 

3. After centrifugation apply the cleared bacterial lysate onto the Glutathione-Sepharose 

column (at 4 ml/min, if using fast-flow material). 

4. After all lysate is applied, wash with standard buffer until the baseline at 280 nm is 

reached again. 

5. Wash with 2 to 4 column volumes standard buffer containing 500 mM KCl and 

1 mM ATP in order to elute nonspecific proteins, especially chaperones that might 

SUPPORT 

PROTOCOL 7 





14.9.17 


SUPPORT 

PROTOCOL 8 


GAP Assays 

14.9.18 

be associated with the desired protein. Afterwards, wash again with standard buffer 

until the baseline at 280 nm is reached. 

High amounts of chaperones indicate folding difficulties with the protein of interest. 

Note that the baseline increases due to 1 mM ATP and drops down after changing to the 

standard buffer with no ATP. 

6. Elute GST-fusion proteins from the column with 100 to 150 ml standard buffer 

containing 20 mM glutathione. Collect fractions of 2- to 5-ml volume using the 

fraction collector provided with the Äkta system. 

The pH value of the standard buffer needs to be readjusted to 7.5 with NaOH after addition 

of glutathione. 

7. Analyze peak fractions by SDS-polyacrylamide gel electrophoresis 

(SDS-PAGE; UNIT 6.1) and Coomassie staining (UNIT 6.6). 

8. Regenerate the column with 50 ml 5 M guanidine hydrochloride and wash with 

100 to 150 ml standard buffer afterwards. 

If the column will not be used for a long period, it should be stored in a buffer containing 

0.01% sodium azide, or, alternatively, in 20% ethanol solution. 

DETERMINING PROTEIN CONCENTRATION USING THE BRADFORD 

ASSAY 

There are different colorimetric and spectrophotometric methods to determine the concentration 

of proteins in a solution including the Lowry, the Smith copper/bicinchoninic 

assay, or the Bradford dye assay. We use both the Bradford dye assay and the Ehresmann 

UV method (Support Protocol 9) for determination of the concentration of a protein in 

solution as well as HPLC in the case of nucleotide binding proteins (also see APPENDIX 3B 

and APPENDIX 3H). 

Materials 

Protein solutions: standards (e.g., BSA or γ-globulin) and test sample 

Bradford reagent (Coomassie dye reagent; Sigma, Pierce, Bio-Rad, or see 

APPENDIX 3H) 

Spectrophotometer 

1. Mix 0.5 ml of Bradford reagent with 1 to 20 μl of a protein solution. For the standard 

curve, mix 0.5 ml of Bradford reagent with 1 to 2 μl of different concentrations of a 

standard protein (e.g., BSA or γ-globulin) covering the range of 0.25 to 2 mg/ml. 

The reagent/protein solution should shift color to blue. 

2. Incubate for 5 min at room temperature. 

3. Measure the absorption of the sample at 595 nm. 

The absorption should be between 0.2 and 0.8 to guarantee correlation between absorption 

and concentration according to the Lambert-Beer law. If the absorption is too low 

or too high, concentrate or dilute the protein solution and reassay. 

4. Determine the protein concentration using the linear absorbance profile of the protein 

standard. 

The Bradford dye solution can be calibrated using different concentrations of a standard 

protein like BSA or γ -globulin. The absorption values of the standards at 595 nm can 

be plotted versus the standard protein concentrations and fit by a linear equation. The 

slope of this regression line will represent a correlation factor between absorbance and 

protein concentration and can in principle be used for the determination of protein 

concentrations. 


DETERMINING PROTEIN CONCENTRATION USING THE EHRESMANN 

ASSAY 

This method is rather useful for small proteins or peptides, since their staining by the 

Bradford dye solution does not lead to a shift of the absorption maxima from 465 nm to 

595 nm comparable to that of BSA or γ-globulin used as standard protein for calibration. 

This method cannot be used for nucleotide binding proteins, since the bound nucleotides 

also absorb at the wavelengths used. 

Materials 

Protein solution 

Quartz cuvettes 

UV/VIS spectrophotometer 

1. Dilute the protein sample to ∼50 μg/ml in deionized water. 

2. Measure the absorption (using a quartz cuvette) of the dilution at 228.5 nm and 

234.5 nm, and use deionized water as a reference. 

3. Calculate the concentration by the following equation: A228.5 − A234.5/3.154 = 

mg/ml. 

No protein standard is used here since the correlation between concentration and absorption 

is due to light absorption of the peptide backbone and not to reactive side chains of 

the proteins. 

CONCENTRATING A DILUTE PROTEIN SOLUTION 

There are a large variety of methods that can be used for concentrating a protein solution, 

including dialysis, ammonium sulfate precipitation (salting out), ion exchange 

followed by desalting chromatography, ultrafiltration or spin-filters, and trichloroacetic 

acid/deoxycholate precipitation. 

We often use the ultrafiltration method by employing spin filters, which accumulate the 

protein at a membrane with a defined molecular weight cutoff (MWCO 5 to 100 kDa) 

while the solvent passes through. 

Materials 

Protein solution 

Refrigerated centrifuge 

Amicon filter, MWCO 5 to 100 kDa 

1. Fill the Amicon device with the diluted protein solution and centrifuge at 2900 × g, 

4 ◦ C. 

2. Continue centrifuging either until a desired concentration is reached (10 to 20 mg/ml 

is a proper concentration for storage) or the volume is sufficient for further purification 

via gel filtration (≤1% of the column volume). 

It is important not to centrifuge to the point where all liquid in the reservoir passes through 

the membrane; this would lead to drying of the protein and consequently to denaturation. 

THROMBIN PROTEOLYTIC CLEAVAGE OF GST FUSION PROTEINS 

The fusion tag that helps to purify a recombinant protein from crude cell extracts should 

be removed when the protein is intended for use in structural or biochemical analysis. 

There are a variety of expression vectors available which have protease-specific cleavage 

sites inserted between the coding sequence for the fusion tag and the multiple cloning site. 

SUPPORT 

PROTOCOL 9 

SUPPORT 

PROTOCOL 10 

SUPPORT 

PROTOCOL 11 





14.9.19 


SUPPORT 

PROTOCOL 12 


GAP Assays 

14.9.20 

The corresponding fusion protein can thus be processed with the appropriate protease 

and, finally, the fusion tag can be removed by further chromatographic purification steps. 

While this protocol is for thrombin cleavage of GST fusion proteins, it can be adapted 

for other enzyme/linker combinations. 

Materials 

GST-fusion protein 

Thrombin (Serva) 

End-over-end rotator 

Additional reagents and equipment for SDS-PAGE (UNIT 6.1) and Coomassie 

staining (UNIT 6.6) 

1. Cleave fusion proteins (at a concentration of ≥1 mg/ml) in batch by incubation with 

1 U thrombin per mg GST-fusion protein for 4 to 16 hr at 4 ◦ C with end-over-end 

rotation. 

Other vectors containing PreScission, factor Xa, TEV, enterokinase, or IgA protease 

cleavage sites can alternatively be used if thrombin does not cleave (masked cleavage 

site) or additional thrombin cleavage sites are within the fusion protein. 

2. Take 10 μl from the reaction mix after 4 hr and after the overnight incubation and 

analyze the cleavage progress by SDS-PAGE (UNIT 6.1) and Coomassie blue staining 

(UNIT 6.6). 

Usually, an overnight incubation at 4 ◦ C is sufficient for a quantitative cleavage of the 

fusion protein. In the SDS-PAGE gel, a band of 26 kDa for GST and a band of the size of 

the fusion partner will appear, whereas the band of the fusion protein at higher molecular 

size will disappear. 

GEL-FILTRATION CHROMATOGRAPHY 

Further purification and removal of protein impurities or small components including 

glutathione are achieved by size-exclusion chromatography (also called gel filtration) on 

the scale of 16/600 or 26/600 columns (meaning columns of 16- or 26-mm diameter and 

600-mm length, respectively) using Superdex 75 or Superdex 200 medium. 

Materials 

Concentrated protein sample (after digestion of the fusion protein with the 

respective protease; see Support Protocol 10 for concentration and Support 

Protocol 11 for digestion) 


16/600 or 26/600 columns prepacked with Superdex 75 or Superdex 200 

(GE Healthcare) 

Äkta System, e.g. Äkta Prime (GE Healthcare) 

Prepacked 16/600 or 26/600 columns with Superdex 75 or Superdex 200 resin (GE 

Healthcare) 

1- to 5-ml sample loop (GE Healthcare) 

Additional reagents and equipment for SDS-PAGE (UNIT 6.1), Coomassie staining 

(UNIT 6.6), and concentration of protein samples (Support Protocol 10) 

1. Equilibrate the column with one column volume (120 ml for 16/600 Superdex or 

320 ml for 26/600 Superdex) of standard buffer using the Äkta System. 

Use a 16/600 column for protein amounts of ≤40 mg or 26/600 column for ≥40 mg but 

≤100 mg; when the protein amount to be purified exceeds 100 mg, divide the sample into 

several portions ≤100 mg and do several consecutive column runs. Use 120 ml of buffer 

for the 16/600 column and 320 ml buffer for the 26/600 column. 


2. Mount a 1- to 5-ml loop (GE Healthcare) on to the Äkta System, flush it with standard 

buffer, and load with the concentrated protein (≥10 mg/ml). 

The volume of the concentrated protein sample to be injected should not exceed 1% of the 

column volume to guarantee efficient separation. In case of a 16/600 column this means 

that no more than 1.2 ml, and in case of a 26/600 no more than 3.6 ml of protein solution 

should be injected. 

3. Collect 1- to 3-ml fractions with the Äkta fraction collector and analyze 10-μl 

samples by SDS-PAGE (UNIT 6.1) and Coomassie staining (UNIT 6.6). 

4. Pool and concentrate the fractions containing the desired protein to 10 to 20 mg/ml 

(Support Protocol 10). 

If the gel filtration does not separate the GST fractions from the desired protein, a 

second affinity chromatography with glutathione-Sepharose is necessary to remove the 

GST completely. 

FREEZING AND THAWING PROTEINS 

Freezing and thawing of protein solutions is a very critical step and has a large impact on 

protein stability. It is absolutely mandatory to freeze a protein solution in liquid nitrogen 

and to store it afterwards at −20◦Coreven−80◦C for storage. For longer storage periods, 

the latter is recommended. Before freezing a protein, try a small-scale test to verify that 

the protein can be frozen in a standard buffer and to test if addition of supplements like 

glycerol or sucrose might help to prevent protein denaturation during freezing. Thawing 

of protein solutions can either be performed rapidly at 37◦C or slowly on ice. For each 

protein, the recommended strategy has to be elucidated in trials. 

Materials 

50- to 500-μl aliquots of purified protein (10 to 20 mg/ml) 


1. Snap freeze 50- to 500-μl aliquots of purified proteins in liquid nitrogen, then store 

in −80 ◦ C freezer. 

2. Thaw all proteins except nucleotide-free GTPases on ice. Thaw nucleotide-free 

GTPases at 37 ◦ C and immediately store on ice when defrosted. 

It is recommended to freeze and thaw an aliquot only once; otherwise, activity of the 

protein might be reduced due to repeated freezing/thawing cycles. Adjust the volume of 

the aliquots to the volume needed for the respective application. However, we have 

observed that most proteins described in this unit (GTPases and GTPase-regulators like 

GEF- and GAP-proteins) can in fact be frozen and thawed at least five times without any 

detectable loss of activity. 




Exchange buffer, 10× 

2M(NH4)2SO4 

10 mM ZnCl2 

Filter and degas 

Store up to several weeks at room temperature 

SUPPORT 

PROTOCOL 13 





14.9.21 



GAP Assays 

14.9.22 

GAP buffer 


10 mM MgCl2 

3 mM dithiothreitol (DTT) 

10 mM potassium phosphate buffer,, pH 7.4 (APPENDIX 2A) 



GEF buffer 


5mMMgCl2 


10 mM potassium phosphate buffer, pH 7.4 (APPENDIX 2A) 



HPLC buffer 

100 mM potassium phosphate buffer, pH 6.5 (APPENDIX 2A) 

10 mM tetrabutylammonium bromide (e.g., Merck) 

7.5% to 25% (v/v) acetonitrile as specified in protocol 



Standard buffer 


5mMMgCl2 


50 mM NaCl 



Terrific broth (TB) medium 

12 g/liter Bacto-tryptone (BD Difco) 

24 g/liter yeast extract (BD Difco) 

0.4% (v/v) glycerol 

2.31 g/liter KH2PO4 

12.54 g/liter K2HPO4 

Sterilize by autoclaving 

Store up to 2 weeks at 4◦C Wash buffer 


5mMMgCl2 


100 mM NaCl 



COMMENTARY 


Fluorescence techniques to study the intrinsic 

properties of GTPase-like nucleotide 

binding, nucleotide exchange, or GTP hydrolysis 

have been evolving over more than two 

decades. They were pioneered by Toshiaki 


Hiratsuka and Roger S. Goody (Alexandrov 

et al., 2001; Hiratsuka, 2003). Since that time, 

further development has been going on, with 

the design of new fluorescent molecules, enhanced 

technical instrumentation, and new applications 

to study biochemical reactions and 

processes. 

Fluorescence-based GEF and GAP assays 

described in Basic Protocols 1 and 2 and 

Alternate Protocols 1 through 4 are convenient 

methods to study the process of nucleotide release 

or hydrolysis. It has been shown that the 

extrinsic fluorophore usually does not influence 

these reactions. Fluorescence-based assays 

allow real-time monitoring of ligand- and 

protein-protein interactions at submicromolar 

concentrations, as well as quantification of the 

kinetic and equilibrium constants. 



Spectroscopic measurements like the fluorescence 

assays described in this unit require 

the use of very clean quartz cuvettes, filtered 

and degassed buffers, and protein solutions 

without precipitate or any other solid material. 

Otherwise, light dispersion will take place 

and the signal-to-noise ratio will deteriorate. It 

is, therefore, very important to centrifuge the 

protein solution immediately before using it in 

assays. 

To obtain reliable and reproducible kinetic 

data, the protein and nucleotide quality need to 

be high. Thus, nucleotides with >90% purity 

should be used and, if necessary, additional purification 

steps should be carried out to obtain 

nucleotides and proteins of a high purity. For 

HPLC measurements, the performance of the 

guard and main columns are essential for an 

optimal separation of the nucleotides. Washing 

the system with filtered and degassed deionized 

water should be carried out daily to prevent 

valve blockage by buffer salts. Regeneration 

of the columns should be accomplished 

periodically according to the column manufacturer’s 

instructions to maintain separation 

performance. 

When purifying guanine nucleotide– 

binding proteins, it is mandatory to add both 

GDP and magnesium ions to the buffer; they 

are essential especially in the case of lowaffinity 

GDP/GTP-binding proteins, and will 

increase the protein stability. 

Releasing the GTPase from GDP (or GTP 

in the case of the constitutive mutants) and 

loading with fluorescent nucleotides, as described 

above, are prerequisites to perform 

fluorescence measurements. The incubation 

time for preparing the nucleotide-free proteins 

varies among GDP/GTP-binding proteins, and 

has to be established for every GTPase. 

The amounts of proteins and (fluorescent) 

nucleotides required are rather dependent on 

the assay used. For the determination of intrinsic 

nucleotide dissociation or intrinsic GTP 

hydrolysis in a cuvette (at a final volume of 

600 μl), ∼10 to 20 μg of nucleotide-bound 

GTPase is required for three identical experiments. 

At least 60 μg of catalytic domains of 

GEF or GAP proteins (250 to 300 amino acids) 

is needed for one experiment to measure the 

specificity and activity of these regulatory proteins. 

A stopped-flow experiment requires 3 to 

5 μg GTPase, but provides an averaged value 

obtained from five to seven identical measurements. 

A stopped-flow analysis of the specificity 

of GEFs or GAPs requires about 15 to 

50 μg of these proteins. However, between 2 

and 10 mg of the catalytic domains will be 

required to quantitatively analyze the GEF or 

GAP activities. 

It is recommended to perform each experiment 

using a negative and a positive control 

sample. The negative sample, necessary 

in order to have correct instrument settings, 

is assessed by measuring free fluorescent nucleotide 

in the absence of GTPase. The positive 

sample, necessary in order to make sure 

that the method works, is assessed by using 

known proteins, for example Ras, to measure 

intrinsic nucleotide dissociation and GTPhydrolysis 

as well as GTPases in combination 

with their specific GEFs (e.g., Ras/Sos1, 

Rac1/Tiam1, Cdc42/Asef or RhoA/p115) or 

GAPs (Ras/p120, Rho GTPases/p50). 


The elucidation of the molecular switch 

mechanism of the GTPases and particularly 

their specificities and affinities for regulators 

require the dissection of such interactions at 

the molecular level by utilizing a sensitive biochemical 

assay. Fluorescence spectroscopic 

methods provide researchers with a number of 

tools for studying the intercommunication of a 

GTPase with nucleotides and binding partners. 

Compared to other qualitative assays (e.g., 

filter-binding assay or thin-layer chromatography, 

which contain between three and six 

time points), fluorescence methods described 

in this unit allow monitoring of the activity 

of the GEFs and GAPs in real time, where 

every single measurement consists of at least 

400 data points per reaction trace. These assays 

are highly sensitive and, in principle, reproducible 

presuming: (1) that the proteins of 





14.9.23 



GAP Assays 

14.9.24 

interest do not reveal the expected activity; 

and (2) that the proteins and reagents are carefully 

prepared from high-purity materials and 

tested for quality. Thus, optimal gene expression 

and protein purification as well as sufficient 

quality of fluorescent nucleotide-bound 

GTPases and other components including GEF 

and GAP proteins and nucleotide derivatives 

are prerequisites for reliable and reproducible 

measurements. In all assays described in this 

unit, a decrease in fluorescence should be monitored 

in a time-dependent manner. However, 

another important aspect to be considered is 

the fluorescence offset, which represents the 

actual fluorescence signal value before the fluorescence 

decay is initiated by starting the reaction. 

This should be relatively similar for all 

experiments (1) when using the same instrument 

settings (e.g. the same monochromator 

slit with in a fluorescence spectrometer and 

photomultiplier tension in case of the stoppedflow 

instrument); (2) under the same concentrations 

of the fluorescent nucleotides in the 

complex with the GTPase; and (3) independent 

of the GEF or GAP concentrations. 

The fluorescence-based assays described in 

this unit are quite sensitive methods which 

give proper results even if using submicromolar 

protein and nucleotide concentrations. The 

fluorescence decay followed over time is usually 

in the range of 40% to 60% in the case of 

nucleotide exchange experiments (GEF assay) 

and 10% to 15% in the case of the fluorescence 

GTP hydrolysis assay using tamra-GTP. 


The expression of a protein in E. coli takes 

about 2 days, including inoculation of the culture 

and induction of gene expression. On the 

second day, the culture is harvested, washed, 

resuspended, and stored in a buffer solution. 

When the expression is carried out for only 

2 to 4 hr, all steps should be performed in 

1 day. The purification of a protein usually involves 

several steps. It takes about 1 day to 

do the affinity chromatography, to elute, to analyze 

by SDS-PAGE, and to concentrate the 

fusion protein before the overnight protease 

digestion. On the next day, the protein can 

be applied to the size-exclusion chromatography 

(gel filtration) and subsequently analyzed 

by SDS-PAGE. Optionally, a second affinity 

chromatography may need to be carried out 

to completely remove the tag; it can be performed 

immediately after gel filtration. Finally 

the protein needs to be concentrated, which 

may take several hours, before it can be frozen 

in small aliquots and stored at –80 ◦ C. 

The preparation of Gpp(NH)p-bound and 

mantGpp(NH)p-bound GTPases takes 2 to 

48 hr, and usually can be carried out overnight. 

The generation of a nucleotide-free GTPase involves 

the same step followed by the phosphodiesterase 

reaction for another 2 to 30 hr until 

Gpp(CH2)p is completely degraded. The loading 

of a nucleotide-free GTPase with mant- 

GDP can be performed in 1 to 2 hr. 

GEF measurements in a fluorescence spectrometer 

usually take at least 16 hr before 

the intrinsic nucleotide dissociation in the absence 

of the GEF protein is recorded. GEFaccelerated 

processes usually take less than 

1 hr. Stopped-flow experiments are performed 

at higher concentrations of the GEF protein 

and are, thus, quite fast reactions on the order 

of seconds or even milliseconds. For quantitative 

measurements and evaluation of the data, 

2 to 3 hr should be allowed. 

Intrinsic GTP-hydrolysis measurements for 

small GTPases take 0.5 to 6 hr (depending on 

the GTPase), whereas the presence of catalytic 

amounts of a GAP protein advances the reaction 

to a 0.5- to 30-min process. Quantitative 

measurements using increasing GAP concentrations 

are also very fast and complete after a 

few seconds or even milliseconds. 


Ahmadian, M.R., Hoffmann, U., Goody, R.S., and 

Wittinghofer, A. 1997a. Individual rate constants 

for the interaction of Ras proteins with 

GTPase-activating proteins determined by fluorescence 

spectroscopy. Biochemistry 36:4535- 

4541. 

Ahmadian, M.R., Stege, P., Scheffzek, K., and 

Wittinghofer, A. 1997b. Confirmation of 

the arginine-finger hypothesis for the GAPstimulated 

GTP-hydrolysis reaction of Ras. Nat. 

Struc. Biol. 4:686-689. 

Ahmadian, M.R., Zor, T., Vogt, D., Kabsch, W., 

Selinger, Z., Wittinghofer, A., and Scheffzek, 

K. 1999. Guanosine triphosphatase stimulation 

of oncogenic Ras mutants. Proc. Natl. Acad. Sci. 

U.S.A. 96:7065-7070. 

Ahmadian, M.R., Wittinghofer, A., and Herrmann, 

C. 2002. Fluorescence methods in the study of 

small GTP-binding proteins. Methods Mol. Biol. 

189:45-63. 

Ahmadian, M.R., Kiel, C., Stege, P., and Scheffzek, 

K. 2003. Structural fingerprints of the Ras- 

GTPase activating proteins neurofibromin and 

p120GAP. J. Mol. Biol. 329:699-710. 

Alexandrov, K., Scheidig, A.J., and Goody, R.S. 

2001. Fluorescence methods for monitoring interactions 

of Rab proteins with nucleotides, Rab 

escort protein, and geranylgeranyltransferase. 

Methods Enzymol. 329:14-31. 

Eberth, A., Dvorsky, R., Becker, C., Beste, 

A., Goody, R.S., and Ahmadian, M.R. 2005. 


Monitoring the real-time kinetics of the hydrolysis 

reaction of guanine nucleotide binding proteins. 

Biol. Chem. 386:1105-1114. 

Guo, Z., Ahmadian, M.R., and Goody, R.S. 2005. 

Guanine nucleotide exchange factors operate 

by a simple allosteric competitive mechanism. 

Biochemistry 44:15423-15429. 

Haeusler, L.C., Blumenstein, L., Stege, P., Dvorsky, 

R., and Ahmadian, M.R. 2003. Comparative 

functional analysis of the Rac GTPases. FEBS 

Lett. 555:556-560. 

Hemsath, L. and Ahmadian, M.R. 2005. Fluorescence 

approaches for monitoring interactions of 

RhoGTPases with nucleotides, regulators and 

effectors. Methods 37:173-182. 

Hiratsuka, T. 2003. Fluorescent and colored trinitrophenylated 

analogs of ATP and GTP. Eur. J. 

Biochem. 270:3479-85. 

John, J., Sohmen, R., Feuerstein, J., Linke, R., 

Wittinghofer, A., and Goody, R.S. 1990. Kinetics 

of interaction with nucleotide-free H-ras 

p21. Biochemistry 29:6058-6065. 

Scheffzek, K. and Ahmadian, M.R. 2005. GTPase 

activating proteins: Structural and functional insights 

18 years after discovery. Cell. Mol. Life 

Sci. 62:3014-3038. 





14.9.25 


In Vivo Imaging of Signal Transduction 

Cascades with Probes Based on Förster 

Resonance Energy Transfer (FRET) 

Takeshi Nakamura 1 and Michiyuki Matsuda 1 

1Laboratory of Bioimaging and Cell Signaling, Graduate School of Biostudies, 

Kyoto University, Kyoto, Japan 

ABSTRACT 

Genetically encoded FRET probes enable us to visualize a variety of signaling events 

such as protein phosphorylation and G-protein activation in living cells. This unit focuses 

on FRET probes wherein both the donor and acceptor are ßuorescence proteins and 

incorporated into a single molecule, i.e., a unimolecular probe. Advantages of these 

probes lie in their easy loading into cells, simple acquisition of FRET images, and 

clear evaluation of data. We have developed FRET probes for Ras-superfamily GTPases, 

designated Ras and interacting protein chimeric unit (Raichu) probes. We hereby describe 

strategies to develop Raichu-type FRET probes, procedures for their characterization, 

and acquisition and processing of images. Although improvements upon FRET probes 

are still based on trial-and-error, we provide practical tips for their optimization and 

brießy discuss the theory and applications of unimolecular FRET probes. Curr. Protoc. 

Cell Biol. 45:14.10.1-14.10.12. C○ 2009 by John Wiley & Sons, Inc. 

Keywords: FRET � unimolecular probe � CFP � YFP � Ras GTPase � Rho GTPase 

INTRODUCTION 

Signal transduction is an organized ensemble of dynamic state changes in signaling 

molecules, which is often caused by phosphorylation and guanine nucleotide exchange 

reactions. Needless to say, the timing and localization of these events critically inßuence 

the outcomes. To visualize such spatio-temporal changes in the activity of signaling 

molecules, several research groups have been developing probes based on the principle 

of Förster resonance energy transfer (FRET; Miyawaki, 2003; Kiyokawa et al., 2006). 

FRET is a radiation-less transfer of excited-state energy from a donor ßuorophore to an 

acceptor ßuorophore (Lakowicz, 2006). This unit focuses on FRET probes wherein both 

the donor and acceptor ßuorophores are proteins and incorporated into a single molecule, 

i.e., genetically encoded unimolecular probes. The advantages of these probes lie in their 

easy loading into cells, simple acquisition of FRET images, and clear evaluation of the 

data (Kurokawa et al., 2004). Another type of FRET probe, namely, the bimolecular 

probe, is preferably used for monitoring protein-protein interactions (UNIT 21.3; also see 

UNITS 17.1 & 17.9). 

A major application of unimolecular FRET probes is the monitoring of protein kinase 

activities. The activities of EGF receptor, Src, PKA, and PKC have been successfully 

monitored by FRET probes containing the speciÞc substrate peptides for each kinase 

(Kurokawa et al., 2001; Ting et al., 2001; Zhang et al., 2001; Violin et al., 2003; Wang 

et al., 2005). Furthermore, FRET probes that monitor the activation-related conformational 

change have also been developed for PKC, Raf, Erk, and CaMKII (Braun et al., 

2005; Takao et al., 2005; Terai and Matsuda, 2005; Fujioka et al., 2006). Visualization 

of the “on” and “off” states of Ras-superfamily GTPases is another valuable application 

of unimolecular FRET probes; such FRET probes developed in the authors’ laboratory 

Current Protocols in Cell Biology 14.10.1-14.10.12, December 2009 

Published online December 2009 in Wiley Interscience (www.interscience.wiley.com). 

DOI: 10.1002/0471143030.cb1410s45 


UNIT 14.10 





14.10.1 

Supplement 45

BASIC 

PROTOCOL 1 

In Vivo Imaging of 



Cascades with 

FRET Probes 

14.10.2 

are collectively designated Ras and interacting protein chimeric unit (Raichu) probes 

(Mochizuki et al., 2001). 

The following procedure describes the development of Raichu-type FRET probes and 

the use of these probes to visualize the activation state of GTPases. 

DEVELOPMENT OF RAICHU FRET PROBES 

Raichu probes are composed of four modules: donor (CFP), acceptor (YFP), a GTPase, 

and the GTPase-binding domain of its binding partner (Fig. 14.10.1). In the Raichu probes 

for Ras-family GTPases, YFP, GTPase, GTPase-binding domain, and CFP are sequentially 

connected from the N-terminus by spacer peptides (Mochizuki et al., 2001). A typical 

mode of action of Raichu is as follows. In the inactive GDP-bound conformation, CFP 

and YFP are located at a distance from each other. Therefore, excitation of CFP results in 

emission mostly from CFP. Upon stimulation, GDP on the GTPase is exchanged for GTP, 

which induces an interaction between the GTP-bound GTPase and the GTPase-binding 

domain. This intramolecular binding brings CFP within close proximity to YFP, thereby 

permitting energy transfer from CFP to YFP. FRET is manifested by a quenching of CFP 

ßuorescence and an increase in sensitized emission of YFP; therefore the ßuorescence intensity 

ratio of YFP versus CFP can be conveniently used as a representation of FRET efÞciency. 

This YFP/CFP value of the Raichu-expressing cells correlates with the molecular 

ratio of GTP-bound probes versus GDP-bound probes, which reßects the balance between 

guanine nucleotide exchange factors (GEFs) and GTPase-activating domains (GAPs) for 

the GTPase within the probe (Mochizuki et al., 2001; Yoshizaki et al., 2003). Therefore, 

the activities of endogenous GTPases, which are under the control of the same set of GEFs 

and GAPs, can be estimated by measuring the YFP/CFP ratio of Raichu-expressing cells. 

YFP 

Ras 

GDP 

activation inactivation 

530 nm FRET 

GTP 

Ras 

433nm 475 nm 

farnesyl moiety 

433 nm 

Supplement 45 Current Protocols in Cell Biology 

YFP 

RBD 

RBD 

Figure 14.10.1 Basic structure of the GFP-based FRET probe for Ras GTPases (Raichu-Ras). 

The FRET probe for Ras activation comprises H-Ras and the Ras-binding domain (RBD) of Raf 

sandwiched between YFP and CFP. For plasma-membrane localization, this probe is fused to the 

carboxyl-terminal region of K-Ras4B. Upon stimulation, GDP on Ras is exchanged for GTP, and 

FRET occurs. 

CFP 

CFP

Materials 

Needed DNA constructs: 

Plasmid for the Raichu probe (available from Matsuda Lab, 

http://www.path1.med.kyoto-u.ac.jp/mm/e-phogemon/index.htm) 

cDNA of the GTPase of interest (can either be purchased from a public 

depository or cloned by PCR from cDNA libraries) 

cDNA of GTPase-binding domains of effector proteins (can either be purchased 

from a public depository or cloned by PCR from cDNA libraries; try at least a 

few known effectors for initial experiments) 

Ion-exchange resin for DNA puriÞcation (e.g., Qiagen; also see Ausubel et al., 

2009) 

Transfection reagent for calcium phosphate coprecipitation (UNIT 20.3) 

293T cells (ATCC cat. no. CRL-11268), cultured in 100-mm-diameter 

collagen-coated dishes, 80% conßuent 

MEM (Invitrogen, cat. no. 10370) containing 10% fetal bovine serum (FBS) 

Phenol-red-free MEM (Invitrogen, cat. no. 11054) containing 10% FBS 

Lysis buffer (see recipe) 

Fluorescence spectrophotometer (for example, JASCO FP-750) and 3-ml cuvettes 

Additional reagents and equipment for basic molecular biology techniques 

including restriction digestion, PCR, plasmid preparation, cloning of DNA, and 

puriÞcation of DNA (Ausubel et al., 2009), and calcium phosphate transfection 

of DNA (UNIT 20.3) 

Make candidate probes 

1. Design a candidate Raichu probe. 

To achieve a wide dynamic range, it is desirable to search for a GTPase-binding domain 

having a moderate afÞnity for the GTPase (Yoshizaki et al., 2003). One explanation for 

this is that the GTPase-binding domain competes with GTPase-activating proteins (GAPs) 

in cells (Kurokawa et al., 2004). This inhibition of GAPs leads to a relatively high GTP 

level in the probe, even in the unstimulated state, and may therefore cause a narrowing 

of the dynamic range. 

Crystallographic data for the GTPase and GTPase-binding domain can help to determine 

the minimum regions to be incorporated into the probe. Unfortunately, crystallographic 

data for optimal design of a Raichu probe cannot currently be obtained in most cases. 

Therefore, various lengths of the GTPase and GTPase-binding domain should be tried 

until a satisfactory result is obtained. In addition, various combinations of the four 

modules, namely, YFP, CFP, GTPase, and the GTPase-binding domain, should be tested. 

YFP is usually placed before CFP because an excess of acceptor (YFP) does not decrease 

the signal-to-noise ratio by much, even when the translation of the probe is prematurely 

terminated. 

Eleven amino acids at the carboxyl terminus of GFP can be truncated without affecting 

its ßuorescence proÞle. In most Raichu probes, we have removed the 11 carboxyl-terminal 

residues of YFP, hoping to reduce the ßexibility between YFP and the subsequent module. 

The length and sequence of the spacers are also critical. If the FRET efÞciency of a prototype 

probe varies to some extent upon activation, the possibility of further improvement 

by changing the spacer should be considered. As spacers, we usually use 1 to 6 repeats 

of the pentapeptide Gly-Gly-Ser-Gly-Gly. It is believed that Gly provides ßexibility while 

Ser prevents aggregation of peptide chains. Misfolding of CFP occasionally occurs, and 

this can sometimes be rectiÞed by modifying the spacer before the CFP. The ideal location 

for a probe in a cell has been a matter of debate. The most persuasive idea is that the 

probe should be colocalized with the endogenous protein; for this purpose, the GTPase’s 

own authentic CAAX-box should be added to the C-terminus of the probe. Alternatively, 

the addition of the CAAX-box of K-Ras4B to the C-terminus enables the probe to be located 

mostly at the plasma membrane; this approach yields a high signal-to-noise ratio, 

especially when only a limited fraction of the GTPase is activated upon stimulation. 





14.10.3 





Cascades with 

FRET Probes 

14.10.4 

2. Construct a plasmid for a candidate probe. 

If you use the basic Raichu probe structure (Fig. 14.10.1), you should Þrst perform PCR 

cloning of the trimmed GTPase and GTPase-binding domain with restriction enzyme 

recognition sites. Thereafter, insert the PCR fragments into the basic FRET vector. 

The 5 ′ and 3 ′ primers for the GTPase should have XhoI and Aor13HI restriction sites, 

respectively. The 5 ′ and 3 ′ primers for the GTPase-binding domain should have KpnI 

and NotI restriction sites, respectively. Any restriction enzymes that generate compatible 

ends can also be used. The primers must be designed so that the inserted cDNA is 

in the correct reading frame of the Raichu (http://www.path1.med.kyoto-u.ac.jp/mm/ephogemon/index.htm). 

The ampliÞed cDNAs of the GTPase and GTPase-binding 

domain are cleaved with the aforementioned restriction enzymes and subcloned into 

XhoI/Aor13HI and KpnI/NotI restriction sites, respectively. The order of the GTPase and 

GTPase-binding domain can be reversed by changing the restriction sites in the primers 

accordingly. 

3. Prepare transfection-quality plasmid DNA. 

DNA puriÞed using Qiagen ion-exchange resin (or equivalents from other manufacturers) 

performs well. Also see Ausubel et al. (2009). 

Characterize candidate probes 

4. Grow 293T cells to 80% conßuence in MEM containing 10% FBS (may contain 

phenol red). Transfect 10 μg of a probe-encoding plasmid into 293T cells using 

calcium phosphate coprecipitation (UNIT 20.3). 

Candidate probes without lipid modiÞcation at the C-terminus are often preferable for 

spectral analysis, because this lipid modiÞcation generally reduces the level of probe 

expression. 

5. At 6 hr after transfection, change the culture medium to phenol red–free MEM/10% 

FBS. 

6. At 48 hr after transfection, harvest the cells in 1 ml lysis buffer. Microcentrifuge the 

lysates at 15 min at 10,000 × g,4 ◦ C. 

The centrifugation serves to clear the lysate. If necessary, the cell lysates may be cleared 

by ultracentrifugation for 15 min at 100,000 × g, 4 ◦ C, to reduce the autoßuorescence 

from cell debris. 

7. Transfer the cleared lysates into 3-ml cuvettes and place the cuvettes in a ßuorescence 

spectrophotometer. Next, illuminate lysates with an excitation wavelength of 433 nm, 

and obtain a ßuorescence spectrum from 450 nm to 550 nm. Subtract the background 

using the spectra of cell lysates prepared without transfection. 

Fluorescence spectral analysis becomes even easier when the FreeStyle 293-F cell line 

(Invitrogen), a variant of the 293 cell line adapted for suspension growth, is used. This 

cell line is very easy to culture, transfect, and harvest, and only 1.5 ml of cell suspension 

is required to obtain a ßuorescence spectrum. 

8. Evaluate a candidate probe using the ßuorescence spectrum (Fig. 14.10.2). 

For the characterization of a candidate probe, we introduce a constitutively active or 

inactive mutation into the GTPase in the probe for comparison with the same probe 

containing the wild-type GTPase. Alternatively, we cotransfect the candidate probe with 

guanine nucleotide exchange factor (GEF) and GAP toward the GTPase, and compare 

the new spectrum with those of samples transfected with the probe alone. Under our 

criteria, Raichu-type probes can be applicable for imaging analysis when the maximum 

increase (%) in the YFP/CFP ratio exceeds 30%. 

Practically, further evaluation of a probe is recommended before use in a wide range of 

applications: i.e., (1) whether the probe shows a near-linear correlation between its GTP 

loading and FRET efÞciency upon cotransfection with various quantities of GEF or GAP 

and (2) whether the probe and its endogenous counterpart show comparable responses 

to physiological stimulations when examined by biochemical methods. 


Emission intensity 

(arbitrary units) 

450 500 

Wavelength (nm) 

Sos (+) 

Sos (-) 

Figure 14.10.2 Fluorescence spectral analysis using 293T cells. Shown are emission spectra 

of Raichu-Ras expressed with the guanidine nucleotide factor Sos (black line, Sos +) or without 

Sos (gray line, Sos −) in 293T cells at an excitation wavelength of 433 nm. 

IMAGING WITH FRET PROBES 

Imaging with FRET probes must be performed with general live-cell imaging precautions, 

and should be modiÞed depending on the speciÞc cell type; these modiÞcations are 

beyond the scope of this unit. Furthermore, the technical details differ among optical 

imaging systems and image analysis software. The following is only a typical example 

of such an analysis. 

Materials 

Cells for experiment [e.g., HeLa cells (ATCC cat. no. CRL-2) or COS7 cells 

(ATCC cat. no. CRK-1651)] and appropriate medium 

Expression plasmid for FRET probe (Basic Protocol 1) 

Transfection reagent 

Phenol red-free medium 

Mineral oil (Sigma) 

35-mm glass-base dish (Asahi Techno Glass; http://www.agc.co.jp/) with a 

10-mm-diameter glass coverslip mounted on the bottom 

Temperature-controlled chamber with thermostat and/or a CO2 controller 

A ßuorescence microscope with a CCD camera including: 

IX81 inverted microscope 

75-W xenon arc lamp 

60× oil immersion objective lens, PlanApo 60×/1.4 (Olympus) 

Cool SNAP-HQ cooled CCD camera (Roper ScientiÞc) 

Laser-based auto-focusing system, IX2-ZDC (Olympus) 

Automatically programmable XY stage, MD-XY30100T-Meta (SIGMA KOKI). 

Excitation and emission Þlter wheels: Filter wheel 99A354 and MAC5000 shutter 

controller (Ludl Electronic Products) 





14.10.5 

Current Protocols in Cell Biology Supplement 45 

550 

BASIC 

PROTOCOL 2




Cascades with 

FRET Probes 

14.10.6 

Filters: 

XF1071 (Omega Optical; cat. no. 440AF21) excitation Þlter 

XF2034 (Omega Optical; cat. no. 455DRLP) dichroic mirror 

XF3075 (Omega Optical; cat. no. 480AF30) emission Þlter for CFP 

XF3079 (Omega Optical; cat. no. 535AF26) emission Þlter for FRET 

Neutral-density (ND) Þlters 

Software for operation of the microscope and analysis of acquired images; e.g., 

MetaMorph software (Universal Imaging) 

Additional reagents and equipment for construction of the FRET probe (Basic 

Protocol 1) 

Prepare cells 

1. Seed the cells onto a glass-base dish with a glass coverslip at the bottom. 

The coverslip at the bottom may be coated with collagen, Þbronectin etc., depending on 

the cell types used. The cell density can be varied depending on the purpose of experiments 

and the transfection protocol. 

2. Transfect cells with the plasmid for a FRET probe using any protocol suitable for 

the cells of interest. 

The transfection efÞciency need not be high, because fewer than 10 cells can be timelapse 

imaged in most experiments. Rather, the transfection method should be chosen to 

minimize cell toxicity. 

3. Grow the cells, typically for 1 to 3 days, depending on the probe and cell type. 

In most cases, within 2 days the amount of probe reaches a level sufÞcient for imaging. 

In cases where the overexpression of the probes is toxic to the cells, the cells should be 

used 1 day after transfection. 

4. On the day of the experiments, change the medium to phenol red-free medium and 

overlay the medium in the cell-containing dish with mineral oil. 

A reduction in the amount of serum is preferable to reduce the background ßuorescence. 

Bovine serum albumin may be included when serum is not included. HEPES should be 

included in the medium unless CO2 is supplied during imaging. In some cases, phosphate 

or HEPES-buffered saline may be preferable to minimize the background ßuorescence. 

Note that some media, such as Opti-MEM, give off high ßuorescence. 

Acquire images of a single Þeld of view 

5. Warm the temperature control chamber to 37 ◦ C and set up ßuorescence microscope. 

A large chamber that covers most of the microscope is preferable to minimize defocusing 

during image acquisition. However, if the microscope is equipped with an autofocusing 

system, the chamber may be as small as needed to keep the cells warm. Always 

use a neutral density (ND) Þlter to minimize the photobleaching of the probe during 

observation. 

6. View the specimen on the microscope using a 60× oil-immersion objective and a 

Þlter set for CFP, and Þnd a cell for time-lapse imaging. 

Cells to be observed should look healthy and represent the cell population. The optimal 

expression level varies depending on each FRET probe. Repeated experiments are required 

to determine the conditions suitable for the imaging. 

7. Set up the image-acquisition software, with the following being a typical setup for 

the FRET imaging of growth factor-induced activation of signaling molecules: 

ND Þlter, 10% 

Camera binning, 4 

Digitizer, 10 MHz 

Gain 2 (4×) 


Shutter speed for phase contrast image or differential interference contrast 

image, 50 msec 

Shutter speed for CFP and FRET, 200 msec 

Time-lapse interval, 1 min 

Number of images, 60. 

The conditions should be optimized for the probes and cells to be analyzed. 

What should be most considered in this process is the photobleaching of YFP and phototoxicity 

to the cells. For an unknown reason, images obtained with shorter exposure time 

and brighter excitation light, which can be easily achieved by removing the ND Þlter, 

usually result in lower FRET efÞciency. This is a principal reason why we prefer the 

conventional epißuorescence microscope to the scanning laser confocal microscope. 

8. Start time-lapse imaging. 

When cells of interest become defocused, stop the image acquisition and focus the cells 

again. Minimize the time to focus, or YFP intensity will be decreased (YFP bleaches faster 

than CFP), resulting in apparent decrease in FRET. 

Acquire images of multiple Þelds of view with autofocusing system 

9. Set up the microscope as described above. Find the Þrst cell to be monitored and 

focus on it. Set the offset value by using the laser-based auto-focusing system. 

Offset value is the distance from the top of the objective lens to the top of the coverslip. 

With a “Þnd focus” command, the autofocusing system remembers the distance between 

the top of the objective lens to the top of the coverslip, and thereby corrects the z-axis 

coordinate prior to image acquisition. Usually, this correction is required only for the 

Þrst cells at each time point. 

10. Find cells to be monitored and record the coordinates. 

The number of cells to be imaged can be reduced afterwards; therefore, a large number 

of cells should be recorded at this stage. When the interval of time lapse is 1 min, 10 to 

15 view Þelds can be imaged. 

11. By moving the motor-driven stage with the obtained coordinates, conÞrm the cells 

to be monitored and correct the focus, if necessary. 

The order of the image acquisition can be changed to minimize the movement of the stage. 

12. Start image acquisition and conÞrm whether a series of image acquisitions Þnishes 

within the interval time. If it goes well, continue image acquisition. 

If not, interrupt image acquisition and reduce the number of cells to be monitored, or 

extend the interval time. It is also strongly recommended to ensure that the hard disc 

contains sufÞcient space for the image. 

Analyze the images 

13. Create stack Þles for the acquired images and overview. 

Carefully examine for defocusing, ßoating bright debris, photobleaching, and phototoxic 

change in the cells. 

14. Create background-subtracted images for CFP and FRET. First, choose region(s) for 

background, where no cells or debris are observed during the entire time-lapse imaging. 

Second, subtract the averaged value of the background region from the image. 

Third, repeat this background subtraction for all planes and make a backgroundsubtracted 

stack Þle. 

The background value should be obtained for each plane. A macro should be prepared 

to repeat this process. Other corrections such as shading can be applied at this stage, if 

necessary. 





14.10.7 





Cascades with 

FRET Probes 

14.10.8 

15. Create ratio images either in simple pseudocolor or intensity-modulated display 

mode. 

The FRET/CFP value is usually used for showing the level of FRET. 

In the intensity-modulated display mode, the brightness and color of each pixel indicates 

the concentration of the probe and the FRET level, respectively. 

16. Create video Þles. From the stacked ratio image, create video Þles either in .avi, 

.mov, or.mpg format. 


Use deionized or distilled water in all recipes and protocol steps. For common stock 

solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX. 

Lysis buffer 


100 mM NaCl 

5mMMgCl2 

0.5% (v/v) Triton X-100 

Store indeÞnitely at 4 ◦ C 

COMMENTARY 


FRET is a process by which a donor ßuorophore 

in an excited state transfers its excitation 

energy to an acceptor nonradiatively 

through dipole-dipole interactions if the emission 

spectrum of the donor overlaps the excitation 

spectrum of the acceptor (Lakowicz, 

2006; UNIT 17.1). This energy transfer manifests 

itself by a quenching of donor ßuorescence 

intensity and lifetime, as well as the 

simultaneous increase in the emission of acceptor 

ßuorescence. Because the efÞciency 

of FRET decreases in proportion to the inverse 

sixth power of the distance between 

the donor and acceptor, FRET has been utilized 

as a spectroscopic ruler to measure 

the distance (

etween two proteins, it is natural to select a bimolecular 

probe (UNIT 17.1 & 21.3). Correction 

of FRET signals obtained with a bimolecular 

probe is elaborate, but executable (Kraynov 

et al., 2000; Sekar and Periasamy, 2003). If 

one wishes to visualize the change of the activity 

of a protein, pH, Ca 2+ concentration, 

etc., a unimolecular probe is preferable; in this 

case, one might take advantage of the merits 

described above. 

Unimolecular FRET probes now cover a 

wide range of signal transduction cascades. 

The archetype is the Ca 2+ sensor, cameleon 

(Miyawaki et al., 1997). The unimolecular 

cameleon has been expressed in speciÞc cell 

types of genetically tractable animals, aiming 

at in vivo Ca 2+ imaging (Kerr et al., 2000; 

Fiala et al., 2002). Concentration of other 

second-messenger molecules such as cyclic 

nucleotides and phosphoinositides can be visualized 

using unimolecular FRET probes 

(Sato et al., 2000, 2003; Zhang et al., 2001). 

Indicators for the on and off states of 

small GTPases such as Ras, Rap1, Ral, R-Ras, 

RhoA, Rac1, and Cdc42 have been developed 

(Mochizuki et al., 2001; Itoh et al., 

2002; Yoshizaki et al., 2003; Takaya et al., 

2004; Pertz et al., 2006; Takaya et al., 2007). 

These FRET probes have been used for the investigation 

of ligand-induced morphological 

change, cell migration, cell division, neurite 

outgrowth, and membrane trafÞcking. FRET 

imaging of Raichu-TC10 by total internal reßection 

microscopy uncovered that the activity 

of TC10 on the exocytic vesicles drops immediately 

before the vesicular fusion (Kawase 

et al., 2006). 

The largest number of unimolecular FRET 

probes are used to monitor protein kinase activities. 

In archetypical probes, an appropriate 

phosphorylation substrate peptide or domain 

and a phosphoamino acid binding domain have 

been linked to produce a hybrid protein, which 

was then ßanked by CFP and YFP. This approach 

has yielded indicators for the activities 

of PKA, PKC, Src, and EGF receptor 

(Kurokawa et al., 2001; Ting et al., 2001; 

Zhang et al., 2001; Violin et al., 2003). In a 

simpler form, the substrate peptide is sandwiched 

between CFP and YFP; conformational 

changes induced by the phosphorylation 

of substrate lead to the change in FRET efÞciency 

(Nagai et al., 2000; Yamada et al., 2005; 

Brumbaugh et al., 2006). Alternatively, entire 

protein kinases are ßanked by YFP and CFP to 

produce a kinase-type probe. Protein kinases 

generally consist of the catalytic and regulatory 

domains, the latter of which associates 

with the former to hold the enzyme in a closed, 

inactive state. Upon stimulation, the regulatory 

domain dissociates from the catalytic domain 

to drive the enzyme into an open, active state. 

This conformational change has been successfully 

monitored as a change in FRET efÞciency 

in the probes for PKC, Raf, Erk, and CaMKII 

(Braun et al., 2005; Terai and Matsuda, 2005; 

Takao et al., 2005; Fujioka et al., 2006). 

Critical Parameters 

For the development of a FRET probe, the 

signal gain, i.e., the difference in the FRET 

ratio (the lowest versus the highest value of 

[YFP ßuorescence]/[CFP ßuorescence] of the 

probe excited for CFP) should exceed 30%. 

If one uses probes in which the gain is less 

than 30%, it is typically difÞcult to observe 

a signiÞcant change in FRET efÞciency upon 

stimulation. 

As for image acquisition, we strongly recommend 

starting with the easiest experiment, 

using established probes, to become familiar 

with the imaging technique. For example, we 

recommend monitoring agonist-induced calcium 

oscillation with a recently developed 

calcium sensor, cameleon YC3.60, which has 

an extraordinarily wide dynamic range (Nagai 

et al., 2004). Imaging of Ras activation in EGFstimulated 

COS cells with Raichu-Ras probe 

(Mochizuki et al., 2001) is another fairly easy 

example (Fig. 14.10.3). 

Recently, the quantitative analysis of acquired 

images has become a necessary skill in 

live cell imaging. Therefore, it is desirable to 

have a good knowledge of image-processing 

software. For that purpose, expert training in 

image processing is helpful. 


Table 14.10.1 provides troubleshooting information 

for these protocols. 


Figure 14.10.3 shows the FRET images of 

COS-1 cells expressing Raichu-Ras following 

EGF stimulation. The FRET images are 

presented in an intensity-modulated display 

mode. EGF-induced Ras activation was detected 

at the periphery of the cell, where membrane 

rufßing was prominent. The increase in 

the YFP/CFP ratio of Raichu-Ras upon EGF 

stimulation is usually 30% to 50%. 





14.10.9 


Table 14.10.1 Troubleshooting Guide to In Vivo Imaging of Signaling Cascades 


Gradual decrease in YFP/CFP 

ratio without stimulation 

Gradual increase in YFP/CFP 

ratio without stimulation 




Cascades with 

FRET Probes 

14.10.10 

Photobleaching during imaging Lengthen time-lapse interval or 

shorten exposure time 

1. Photobleaching during 

searching for the target cell. 

2. Progress of folding of 

ßuorescence proteins. In this 

case, both YFP and CFP 

intensities should increase. 

Defocus during imaging Distortion of warmed 

microscope 

Higher (or lower) YFP/CFP ratio 

in speciÞc side of the image 

Appearance of paired high and 

low areas of YFP/CFP ratio at 

the edge of cells 

Cells expressing FRET probes look 

unhealthy 

Wait for 5-10 min and restart the 

imaging. 

Prolong the incubation time after 

probe transfection. 

Warm up microscope and chambers 

in advance. 

To avoid defocus completely, a 

focus-drift compensation system (e.g., 

Olympus IX2-ZDC) is very useful. 

Uneven illumination to samples Align an arc lamp precisely 

Misregistration (subtle 

displacement between CFP 

and YFP images) 

Cell toxicity due to the 

overexpression of probes 

Correct misregistration using 

imaging software 

Choose the healthy cells expressing 

lowlevelsofprobesorretrythe 

transfection with less DNA 

Figure 14.10.3 Time-lapse experiment of Ras activation upon EGF stimulation in COS-1 cells. 

Serum-starved COS-1 cells expressing Raichu-Ras were treated with 50 ng/ml of EGF and imaged 

every 2 min. FRET images are shown at the indicated time points. For color figure go to 

http://www.currentprotocols.com/protocol/cb1410. 


In some cases, more than one hundred candidate 

probes should be designed, constructed, 

and checked for FRET efÞciency to obtain an 

ideal probe. Thus, the term needed to develop 

a good probe is ∼3 months. This time frame 

signiÞcantly varies, depending on the knowledge 

of the target for probe development. 

The entire cell-imaging procedure can be 

performed in 3 to 4 days from cell seeding. 

This period includes 1 day for expression of 

FRET probe after transfection, 1 day for data 

acquisition, and 1 day for data analysis. 


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14.10.11 





Cascades with 

FRET Probes 

14.10.12 

Wang, Y., Botvinick, E.L., Zhao, Y., Berns, M.W., 

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Key References 

Miyawaki, 2003. See above. 

Comprehensive review of GFP-based FRET technology. 

Internet Resources 

http://www.path1.med.kyoto-u.ac.jp/mm/ 

e-phogemon/index.htm 

Web site for further information about the Raichutype 

FRET probe. Setup of the FRET imaging system 

is described.

"Signal Transduction: Protein Phosphorylation". In: Current Protocols ...

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