PhD Thesis - Biologisk Institut

F A C U L T Y O F S C I E N C E
U N I V E R S I T Y O F C O P E N H A G E N
PhD Thesis
Sine Godiksen
Towards an Understanding of the Role of
Matriptase in Normal Physiology
Academic advisors: Lone Rønnov-Jessen and Lotte K. Vogel
Submitted: 17/09/2012

Towards an Understanding of the Role of
Matriptase in Normal Physiology
Submitted to:
The Graduate School of Science, Faculty of Science
Department of Biology, University of Copenhagen, Denmark
For the PhD Degree
by
Sine Godiksen
Department of Biology, University of Copenhagen
Universitetsparken 13, 2100 København Ø
and
Department of Cellular and Molecular Medicine, University of Copenhagen
Blegdamsvej 3, 2200 København N
September 2012

Preface
This thesis is submitted to the Faculty of Science, University of Copenhagen, Denmark as basis for
obtaining the PhD degree. I received a personal scholarship from the Faculty of Science in October
2007 and started my PhD studies in January 2008. My PhD thesis entitled: ”Towards an
understanding of the role of matriptase in normal physiology” is based on the work performed
from November 2009 to August 2011 in the Lab of Lotte Vogel, Department of Cellular and
Molecular Medicine, Faculty of Health and Medical Sciences, University of Copenhagen, with Dr.
scient Lone Rønnov-Jessen, Department of Biology, Faculty of Science, University of Copenhagen
as faculty supervisor and Lotte Vogel as supervisor.
Additionally, I have included work performed in the Lab of PhD Thomas H. Bugge, National
Institutes of Health, Bethesda, MD, USA under a leave of absence from my PhD studies from
September 2011 – April 2012 (paper II).
The data obtained for this thesis have been presented both at national and international scientific
meetings and the results have resulted in three scientific papers; two published manuscripts and a
manuscript in progress (see list of papers).
My research was financially supported by the Faculty of Science, Danish Cancer Society, the
Augustinus Foundation, Købmand Kristian Kjær og Hustrus Foundation - the Kjær-Foundation,
Dagmar Marshall´s Foundation, Snedkermester Sophus Jacobsen og Hustru Astrid Jacobsen´s
Foundation, Grosserer Valdemar Foersom og Hustru Thyra Foersom´s Foundation, Fabrikant Einar
Willumsens Mindelegat.
Sine Godiksen
Cand. Scient.
Copenhagen, September 2012

Table of Contents
ACKNOWLEDGEMENTS .......................................................................................................... 3
LIST OF PAPERS ...................................................................................................................... 4
ABSTRACT ............................................................................................................................... 5
DANSK RESUMÉ ...................................................................................................................... 7
LIST OF ABBREVIATIONS ....................................................................................................... 9
INTRODUCTION ...................................................................................................................... 11
AIMS AND OBJECTIVES ........................................................................................................ 13
BACKGROUND ....................................................................................................................... 14
Membrane-anchored serine proteases ................................................................................................................14
Matriptase ...........................................................................................................................................................16
Matriptase expression and structure .......................................................................................................................... 16
Regulation of matriptase ......................................................................................................................................18
Matriptase inhibitors .................................................................................................................................................. 18
Matriptase proteolytic processing and activation ...................................................................................................... 20
Prostasin ..............................................................................................................................................................22
Physiological functions of matriptase and its role in pathological processes ........................................................23
Matriptase is crucial for epidermal integrity .............................................................................................................. 23
The matriptase-prostasin proteolytic pathway(s) in the epidermis ........................................................................... 24
Matriptase has an important role in epithelial homeostasis ...................................................................................... 25
Importance of matriptase regulation in vivo .............................................................................................................. 26
Matriptase in carcinogenesis ...................................................................................................................................... 27
TECHNICAL CONSIDERATIONS............................................................................................ 29
Cell system ...........................................................................................................................................................29
Protease pull down assays ...................................................................................................................................30
Biotinylation assays..............................................................................................................................................32
RESULTS ................................................................................................................................ 35

Paper I ..................................................................................................................................................................36
Paper II .................................................................................................................................................................47
Paper III ................................................................................................................................................................65
DISCUSSION AND PERSPECTIVES ...................................................................................... 85
REFERENCES ......................................................................................................................... 91
SUPPLEMENTARY I ............................................................................................................. 105
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Acknowledgements
During the process of gaining experimental data and knowledge to complete this thesis and
obtain my PhD there are many people whom I would like to thank. Without their continuous help,
encouragement and support, this work would not have been possible. I would hereby like to
express my gratitude to:
Lone Rønnov-Jessen for taking on the job as my supervisor in difficult times. Thank you for your
great support and for your invaluable guidance in my writing process.
Lotte K. Vogel for giving me the opportunity to finish my PhD studies, and for your support
throughout my PhD.
Thomas Bugge for giving me a very interesting and inspiring stay in his lab in Bethesda and for his
excellent scientific guidance.
Jan K. Jensen for valuable discussions throughout my PhD studies.
All of my colleagues at ICMM, University of Copenhagen and NIDCR, National Institutes of Health
(NIH), for a very pleasant, helpful and inspiring environment.
Stine in particular, for support, scientific discussions, and for establishing the getaway to NIH.
Karen Skriver for your interest and deeply valued support throughout my PhD.
Annette Storgaard for your professionalism and for your guidance.
Family and friends for your constant support and encouragements.
And finally my beloved Peter, “tak for mad” and for being the most supportive and understanding
“kæreste” anyone could ask for.
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List of papers
Papers included in the thesis:
Paper I
Transport via the transcytotic pathway makes prostasin available as substrate for matriptase.
Stine Friis, Sine Godiksen, Jette Bornholdt, Joanna Selzer-Plon, Hanne Borger Rasmussen, Thomas
H. Bugge, Chen‐Yong Lin and Lotte K. Vogel
Paper II
Reduced prostasin (CAP1/PRSS8) activity eliminates HAI-1 and HAI-2 deficiency-associated
developmental defects by preventing matriptase activation. Roman Szabo, Katiuchia Uzzun
Sales, Peter Kosa, Natalia A. Shylo, Sine Godiksen, Karina K. Hansen, Stine Friis, Silvio Gutkind,
Lotte K. Vogel, Edith Hummler, Eric Camerer, and Thomas H. Bugge
Paper III
Novel assay for detection of active matriptase.
Sine Godiksen and Lotte Vogel. Other authors to be included.
Papers not included in the thesis:
Bornholdt J, Friis S, Godiksen S, Poulsen SS, Santoni‐Rugiu E, Bisgaard HC, Bowitz Lothe IM, Ikdahl
T, Tveit KM, Johnson E, Kure E, Vogel LK. The level of claudin‐7 is reduced as an early event in
colorectal carcinogenesis. BMC Cancer, 2011, Feb 10; 11:65.
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Abstract
Matriptase is a type II membrane-anchored serine protease essential for epithelial integrity and
with implication in carcinogenesis. Matriptase is co-expressed in epithelial cells of most tissues
with the protease prostasin and their mutual inhibitor hepatocyte growth factor activator
inhibitor 1 (HAI-1). All three proteins have crucial functions for epidermal integrity; and in the
epidermis matriptase acts upstream of prostasin and is required for its activation.
The difference between the subcellular locations of matriptase and prostasin has so far been an
enigma for matriptase-prostasin interaction. By mapping the subcellular trafficking of matriptase,
prostasin and HAI-1, we demonstrate that the basolateral plasma membrane is part of the
subcellular itinerary of both proteases and thus constitutes a location for interaction. In Caco-2
cells, zymogen matriptase is routed to the basolateral plasma membrane where it becomes
activation site cleaved and activated. At steady state, prostasin is found to locate mainly to the
apical plasma membrane although a minor fraction of prostasin can be detected at the
basolateral plasma membrane. We show that prostasin is present in its active form on both the
apical and basolateral plasma membrane and that prostasin as well as HAI-1 are endocytosed
from the basolateral plasma membrane and transcytosed to the apical plasma membrane.
Activation of matriptase on the basolateral plasma membrane is followed by efficient
internalization as a matriptase-HAI-1 complex.
Deregulated or unopposed matriptase activity causes a perturbation of several biological
functions e.g. developmental defects in mice. However, the molecular mechanisms behind these
effects are poorly understood. We performed a genetic epistatic analysis to identify new
components of matriptase-dependent pathways in embryogenesis. We show that prostasin is an
indispensable component of the matriptase-dependent proteolytic cascade that causes early
embryonic lethality in mice. In placental tissue prostasin acts upstream of matriptase and is
required for the activation of matriptase. During embryonic development both HAI-1 and HAI-2
are essential inhibitors of matriptase. In this study we moreover find that both HAI-1 and HAI-2
are indispensable inhibitors of prostasin in placental tissue.
Matriptase activity is a tightly regulated protease; zymogen and HAI-1-complexed matriptase is
abundantly present in most epithelial cell lines and tissues. There are no specific substrates or
inhibitors of matriptase, which has led to challenges in detecting active matriptase. In order to
assess the location and relative amount of active matriptase as compared to the total amount, we
have established an assay for detection of active matriptase. This assay is based on a
chloromethyl ketone peptide inhibitor with a predicted substrate sequence of matriptase. We
show that active matriptase is present on the basolateral plasma membrane of Caco-2 cells and
merely comprise a fraction of total matriptase. The conversion of zymogen matriptase to active
matriptase can be induced by exposure to slight acidic conditions as well as the levels of HAI-1-
complexed matriptase and active matriptase rises under these conditions in Caco-2 cells. Labeling
of Caco-2 cells with the chloromethyl ketone peptide inhibitor delayed matriptase-HAI-1 complex
formation. In addition to the peptide inhibitor being unable to bind HAI-1-complexed matriptase,
we show that the matriptase zymogen has an intrinsic activity that enables it to bind the
chloromethyl ketone peptide inhibitor. Moreover, this assay can be easily modified to detect
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active matriptase in other cell system, and we show that cultured primary murine keratinocytes
contain low levels of peptidiolytic active matriptase.
Taken together, these results have improved our knowledge of the interplay between matriptase
and prostasin towards a understanding of these components roles in epithelial integrity and
present an assay for detection of active matriptase.
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Dansk resumé
Matriptase er en type II membranbunden serinprotease, der er essentiel for integriteten af
epitelvæv og er endvidere involveret i udvikling af kræft. Matriptase ekspreseres sammen med
proteasen prostasin og deres fælles proteasehæmmer hepatocyte growth factor activator
inhibitor 1 (HAI-1) af epitelceller i mange væv. Alle tre proteiner har en vital funktion for
integriteten af epidermis, og matriptase er nødvendig for aktivering af prostasin i dette væv.
Forskelle i subcellulær lokalisering mellem matriptase og HAI-1 på basolateralmembranen og
prostasin på apikalmembranen af polariserede epitelceller har hidtil været et uafklaret dilemma i
forhold til de to proteasers samspil. Kortlægning af den intracellulære transport af matriptase,
prostasin og HAI-1 i Caco-2 celler angiver den basolaterale plasmamembran som en mulig
subcellulær lokalitet, hvor matriptase og prostasin kan interagere. Zymogen matriptase
exocyteres til basolateralmembranen, hvor proteasen kløves proteolytisk til den aktive form. Ved
”steady state” befinder prostasin sig primært på apikalmembranen, omend en mindre fraktion
kan detekteres på basolateralmembranen. Vi viser, at aktivt prostasin findes på både apikal -og
basolateralmembranen, og at prostasin såvel som HAI-1 internaliseres fra basolateralmembranen
og transcyteres til apikalmembranen. Aktivering af matriptase på basolateralmembranen
efterfølges af internalisering af matriptase i kompleks med HAI-1.
Dereguleret og ukontrolleret matriptase aktivitet er en tilgrundliggende årsag til uligevægt i flere
vigtige biologiske processer, f.eks. udviklingsmæssige defekter i mus.
Vi udførte genetiske analyser i mus for at identificere nye komponenter i matriptase-afhængige
signalveje. Disse resultater viser, at prostasin er en essentiel komponent i den matriptaseafhængige
proteolytiske kaskade, der resulterer i embryonal dødelighed i mus. Vi viser, at
prostasin ligger opstrøms for matriptase og er nødvendig for aktivering af matriptase i murinet
placental væv. Både HAI-1 og HAI-2 er tidligere vist at være vigtige proteasehæmmere af
matriptase i placenta. I dette studie viser vi, at dette også er gældende for prostasin.
Matriptase er højt reguleret på posttranslationel niveau, hvilket indikeres af, at zymogen
matriptase og matriptase i kompleks med HAI-1 let detekteres i mange cellelinier og epitelvæv.
Der er ikke identificeret en specifik inhibitor eller et specifikt substrate for matriptase, hvilket har
betydet, at detektering af aktivt matriptase har været udfordrende.
Vi udviklede et assay til detektering af aktivt matriptase baseret på en klorometylketon
peptidinhibitor med en peptidsekvens, der afspejler en foretruknen substratesekvens for
matriptase.
Vi viser, at aktivt matriptase findes på basolateralmembranen; denne udgør dog kun en fraktion af
total matriptase. Zymogen aktivering af matriptase kan induceres ved et let fald i pH, hvormed
mængden af aktiv matriptase stiger, ligesom kompleksdannelse med HAI-1 induceres under disse
forhold. Ydermere forsinker mærkning med klorometylketon peptidinhibitoren kompleksdannelse
mellem matriptase og HAI-1; tilsvarende er inhibitoren ikke i stand til at mærke matriptase-HAI-1
komplekset. Vi viser endvidere, at zymogen matriptase er i stand til at binde klorometylketone
peptidinhibitoren. Ydermere kan dette assay let modificeres til andre cellesystemer, og vi viser
med dette assay tilstedeværelsen af aktivt matriptase i kulturer af primære murine keratinocytter.
7

Disse resultater har udvidet vores viden om samspillet mellem matriptase and prostasin mod en
forståelse af disse proteases rolle for integriteten af epitel og præsenterer et assay til detektering
af aktiv matriptase.
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List of Abbreviations
aa
Amino acid
Arg
Arginine
ARIH
Autosomal recessive ichthyosis with hypotrichosis
Asp
Aspartic acid
cDNA
Copy deoxyribonucleic acid
CAP
Channel activating protease
CDCP1 CUB domain-containing protein 1
CUB
Cls/Clr, urchin embryonic growth factor, and bone morphogenic protein-1
DESC
Differentially expressed in squamous cell carcinoma gene
EGFR
Epidermal growth factor receptor
ENaC
Epithelial Na + channel
ER
Endoplasmic reticulum
Gly
Glycine
GPI
Glycosylphosphatidylinositol
Gpld1
GPI-specific phospholipase D1
HAI-1 Hepatocyte growth factor activator inhibitor 1
HAI-2
Hepatocyte growth factor activator inhibitor2
HAT
Human airway trypsin-like
HGF/SF
Hepatocyte growth factor/scatter factor
HGFA
Hepatocyte growth factor activator
His
Histidine
IGFBP-rP1
Insulin-like growth factor binding protein-related protein1
IP
Immunoprecipitation
KD1
N-terminal Kunitz domain
KD2
C-terminal Kunitz domain
kDa
Kilo dalton
LDLR
Low density lipoprotein receptor class A
Lys
Lysine
MAM
Meprin, A5 antigen, and receptor protein phosphatase μ
MANSC
Motif at the N-terminal containing seven cysteines
MDCK
Mardin Darby canine kidney (cell)
mRNA
Messenger ribonucleic acid
MMP-3 Matrix metalloproteinases 3
MSP-1
Macrophage stimulating protein-1
MSPL
Mosaic serine protease large-form
MT-SP1
Membrane type serine protease 1 (matriptase)
NHS
N-hydroxysulfosuccinimide
PAR-2
Protease-activated receptor-2
PDGF
Platelet-derived growth factor
PN1 Protease nexin 1
9

PRSS14 Protease serine S1 family member 14
PS-SCL
Positional scanning synthetic combinatorial library
S1P
Sphingosine 1-phosphate
SA
Signal anchor
SDS-PAGE
Sodium dodecyl sulphate polyacrylamide gel electrophoresis
SEA
Sea urchin sperm protein, enteropeptidase, and agrin
Ser
Serine
SIMA135
Subtractive immunization M(+)HEp3 associated 135 kDa protein
S-NHS-SS-Biotin Sulfosuccinimidyl-2-[biotinamido]ethyl-1,3-dithiopropionate
SPD
Serine protease domain
ST14
Suppressor of tumorigenicity-14
TADG-15
Tumor-associated differentially expressed gene-15
TEER
Transepithelial electrical resistance
TMPRSS
Transmembrane protease, serine
tPA
Tissue plasminogen activator
TTSP
Type II transmembrane serine protease
uPA
Urokinase-type plasminogen activator
VEGFR-2 Vascular endothelial growth factor receptor 2
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Introduction
Proteolysis is a critical event in many biological processes from simple protein degradation in
nutrient digestion to specific alterations in protein activity in highly regulated protease cascades,
e.g. the blood coagulation system [1]. Formerly serine proteases represented enzymes that were
either secreted or sequestered in cytoplasmic storage organelles awaiting signal-regulated
release, but analysis of the human genome at the turn of the millennium revealed a new subclass
of serine protease; the membrane-anchored serine proteases. With this family, a whole new
range of important biological processes was shown to be regulated by serine proteases [2;3].
Matriptase is a type II membrane-anchored serine protease and is especially interesting due to its
role in carcinogenesis and due to its importance for development and maintenance of epithelial
integrity [4-8]. However, little is known of matriptase´s role at the molecular level in
carcinogenesis. Of importance is the finding that a modest over-expression of wild-type
matriptase in the epidermis of transgenic mice is sufficient to promote spontaneous squamous
cell carcinoma formation. A simultaneous increase in expression of matriptase´s cognate inhibitor
hepatocyte growth factor activator inhibitor 1 (HAI-1) completely negates the oncogenic
phenotype of matriptase over-expression suggesting that the unopposed matriptase activity is the
underlying cause [6].
Studies of matriptase knock-out mice have revealed critical functions for matriptase in
development and maintenance of multiple epithelial tissues, which is in accordance to its
widespread epithelial distribution in both mice and humans [4;5;9-12]. In humans, mutations in
the ST14 gene encoding matriptase cause congenital ichthyosis [13-15]. Matriptase acts upstream
of the glycosylphosphatidylinositol (GPI)-anchored serine protease prostasin in terminal
epidermal differentiation and evidence suggests that an impairment of this cascade is the
underlying cause of the symptoms observed in these patients [13;15-19]. Moreover, inhibition of
matriptase is an essential function of HAI-1 in maintaining epidermal homeostasis in mice and
zebrafish, and HAI-1 has been shown to inhibit both matriptase and prostasin in cultured human
keratinocytes [20-22].
In contrast to most enzymatic reactions, proteolysis is a “one-way” process, which requires strict
regulation of protease activity. Spatial distribution and confinement of proteases to specific
cellular compartments is one way to ensure that proteolytic processing occurs under the correct
conditions. Matriptase, prostasin and HAI-1 are co-expressed in polarized epithelial cells and HAI-
1 is an inhibitor of both proteases [9;11;21;23;24]. Polarized eepithelial cells have a plasma
membrane that is divided into an apical compartment and a basolateral compartment. Both HAI-1
and matriptase is located at the basolateral plasma membrane in numerous epithelial tissues and
cell types whereas, prostasin is located at the apical plasma membrane [11;23-26]. This difference
in spatial distribution is in conflict with the proposed matriptase-prostasin proteolytic cascade
[16].
We have earlier shown in a recombinant system that HAI-1 is transported to the basolateral
plasma membrane. After endocytosis, a fraction of HAI-1 is transcytosed across the polarized cell
to the apical membrane compartment, suggesting how HAI-1 can access and inhibit both
matriptase and prostasin (supplementary I; [27]). However, it is still uncertain how matriptase is
11

able to activate prostasin in light of their different subcellular localization. Nevertheless, the
global co-expression of matriptase and prostasin could indicate that this proteases cascade has a
general role in maintaining epithelial integrity. Still, it remains unclear what substrates and
pathways are involved in matriptase-dependent epithelial homeostasis, and if prostasin is a global
downstream target for matriptase.
The life cycle of matriptase is very complex and tightly regulated. Matriptase is believed to have a
role as a protease at the pinnacle of protease cascades due to its ability to autoactivate [28;29].
Matriptase activation and inhibition by HAI-1 is shown to be intimately linked and has lead to the
hypothesis that matriptase only exist in its free active form for a limited period and therefore only
has a narrow time window to act on its substrate(s) before being pacified by inhibitor complex
formation [21;30;31]. Thus, experimental tools for detection of active matriptase are highly
desirable.
Hence, matriptase is an important serine protease essential for epithelial integrity and with
implications in cancinogenesis. Still, it remains to be elucidated where in the polarized cell
matriptase is activated and able to act on downstream substrate(s). Moreover, identification of
additional components of matriptase-dependent proteolytic pathways would provide valuable
insights into the biology of matriptase-dependent biological processes.
This thesis is written as a synopsis and gives a brief introduction to membrane-anchored serine
proteases and matriptase in particular. Mechanisms for matriptase activation and inhibition will
be described with an introduction of matriptase´s most important inhibitors, HAI-1 and HAI-2, and
its epidermal downstream target prostasin. Finally, a description of matriptase in relation to its
physiological functions and its role in pathological processes will be given.
The background section is followed by aims and objectives of this thesis and technical
considerations. The results obtained are enclosed as two published papers and one manuscript in
progress. Finally, the results will be discussed in relation to previous studies.
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Aims and objectives
The overall aim of this study was to gain a more comprehensive understanding of matriptase in
epithelial biology.
We wanted to delineate how matriptase can act as an upstream activator of prostasin in
epidermal terminal differentiation, despite their different subcellular localization in polarized
epithelial cells at steady state, by mapping the intracellular trafficking of matriptase, prostasin and
HAI-1 (paper I). We furthermore wished to determine where in the polarized cell matriptase is
activated and able to cleave its substrates (paper I and paper III). More so, we wanted to establish
an assay for detection of active matriptase on the surface of living cells (paper III). One way of
elucidating the physiological functions of proteins is by identification of interaction partners,
upstream regulators and downstream effectors. Accordingly, we aimed at identifying new
components of matriptase-dependent proteolytic pathways critical for embryogenesis in mice
(paper II).
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Background
To understand the importance of matriptase and the relationship between matriptase, prostasin
and their inhibitor HAI-1, this chapter gives an introduction to these proteins as well as the
mechanisms of matriptase activation. Afterwards, I will outline important physiological functions
of matriptase, its interplay with its inhibitors and prostasin in these processes, and finally review
the role of matriptase in cancer.
Membrane-anchored serine proteases
Proteases are enzymes that conduct proteolysis and were originally discovered a century ago as
enzymes involved in nutrient digestion. Sequencing of the human genome at the turn of the
millennium revealed a total 569 proteases in humans. On the basis of the mechanism of catalysis,
proteases are classified into five distinct classes: aspartic, metallo, cysteine, serine and threonine
proteases [32].
Serine proteases are one of the largest and most conserved multigene proteolytic families. This
protease family includes the well-known digestive enzymes trypsin and chymotrypsin. Serine
proteases are found in a variety of tissues and body fluids with well characterized roles in diverse
cellular functions including blood coagulation, wound healing, digestion and immune response.
Furthermore, many studies indicate that these proteases contribute to a number of pathological
conditions and play a significant role in tumour growth, invasion and metastasis [33;34].
Serine proteases are characterized by the presence of a serine residue in the active site of the
catalytic domain. In the active site resides the catalytic triad that is preserved in all serine
proteases. This catalytic triad is composed of three amino acids; a histidine, a serine and an
aspartic acid, which are essential for the catalytic ability of the protease. The specificity of the
protease is determined by the size, shape and charge of the active site cleft [33].
The rate of discovery of new (serine) proteases was greatly aided and accelerated by the
publishing of the mouse and human genome sequences and EST databases. This led to the
discovery of the hitherto unknown large family of membrane-associated serine proteases; the
membrane-anchored serine proteases. Orthologues are found in all vertebrates and in humans 20
members have been identified so far as presented in fig. 1 [35;36].
Membrane-anchored serine proteases are tethered to the membrane either via a C-terminal
transmembrane domains (Type I), a GPI-anchor or via an N-terminal transmembrane domain
(Type II) as illustrated in fig. 1. The common features of Type II transmembrane serine proteases
(TTSP) are a short cytoplasmic anchor, an N-terminal transmembrane domain, a C-terminal
extracellular serine protease domain of the trypsin (S1) fold, and a variable length stem region
containing modular domains linking the catalytic and the transmembrane domains. This stem
region contains an assortment of 1-11 protein domains of six different types [3;35].
14

Fig 1. Overview and domain structure of the human membrane-anchored serine proteases.
Prostasin and testisin is anchored to the membrane by a GPI-anchor, whereas tryptase γ1 is
anchored to the membrane through a C-terminal transmembrane domain (type I transmembrane
protein). The remaining membrane-anchored serine proteases presented here are type II
transmembrane proteins and are attached to the membrane by a signal anchor (SA) located close
to the N terminus with a cytoplasmic extension. Type II transmembrane serine proteases are
phylogenetically divided into four subfamilies; the largest being the human airway trypsin-like
(HAT)/differentially expressed in squamous cell carcinoma gene (DESC) subfamily, which comprises
HAT, DESC1, TMPRSS11A, HAT-like 4, and HAT-like 5 (blue shading); the hepsin/transmembrane
protease, serine (TMPRSS) subfamily, which comprises hepsin, TMPRSS2, TMPRSS3, TMPRSS4,
mosaic serine protease large-form (MSPL), spinesin, and enteropeptidase (yellow shading); the
matriptase subfamily, which consists of matriptase, matriptase-2, matriptase-3, and polyserase-1
(green shading); and the corin subfamily, which contains only corin (purple shading). Beside a
serine protease domain (SPD) of the S1 fold, the modular structure of the TTSP can contain an
assortment of different domains that include sea urchin sperm protein, enteropeptidase, and agrin
(SEA); group A scavenger receptor (scavenger); low-density lipoprotein receptor class A (LDLR);
Cls/Clr, urchin embryonic growth factor, and bone morphogenic protein-1 (CUB); meprin, A5
antigen, and receptor protein phosphatase μ (MAM); and frizzled. The figure is from [3].
15

This complex modular structure of most TTSPs is remarkably different from other trypsin-like
serine proteases. The multi-domain structure provides the TTSPs with the capacity to interact
with several interaction partners and possibly a mean for regulation of proteolytic activity [36;37].
Several TTSPs are involved in fundamental cellular and developmental processes such as
morphogenesis, differentiation, epithelial permeability and cellular iron transport [3].
Matriptase
Matiptase was first identified in 1993 as a new gelatinolytic activity expressed by cultured breast
cancer cells and later 5 different groups individually cloned the cDNA from human prostate cancer
cells, human ovarian carcinoma, human breast cancer cells, murine thymic cells, and human colon
mucosa [38-43]. Shortly after matriptase was purified from human milk [44]. The molecular
cloning of the two closely related proteases matriptase-2 and matriptase-3 was reported later
[45;46]. Matriptase is also referred to as MT-SP1, TADG-15, PRSS14, TMPRSS14, SNC19, cleaving
activating protease (CAP)-3, prostamin, epithin (mouse ortholog) and the gene designation is
suppression of tumorigenicity 14 (ST14). For simplicity, I will use matriptase also when discussing
the mouse orthologue.
Matriptase expression and structure
Orthologues of matriptase has been found in many vertebrates but unlike most other TTSPs,
matriptase is widely expressed in epithelial compartments of many embryonic and adult tissues
[10;11]. Expression analysis has been done on both human and murine tissues. mRNA and protein
analyses show that matriptase is expressed in all types of epithelium, including columnar,
pseudostratified, cubiodal, and squamous in humans and locates to the basolateral plasma
membrane [11]. Using enzymatic gene trapping with -galactosidase as a reporter in mice showed
an expression pattern of matriptase in mice overall identical to what was found in humans [12].
The high degree of similarity between expression profiles of different species suggests conserved
function(s) of matriptase.
Full-length human matriptase is an 855 amino acid (aa) glycoprotein that lacks a classical signal
peptide of app. 95 kDa. Instead the N-terminal signal anchor, which is not removed during
synthesis, functions as a single-span transmembrane domain that orientates the protease in the
plasma membrane as a type II integral membrane protein (fig. 1 and 2). Matriptase is a mosaic
protein and is composed of a short N-terminal tail (residues 1-54) followed by the transmembrane
domain, a sea urchin sperm protein, enterokinase, agrin (SEA) domain (residues 85-193), two
C1r/s, urchin embryonic growth factor and bone morphogenetic protein 1 (CUB) domains
(residues 214-334 and 340-447), four low-density lipoprotein receptor class A (LDLR) domains
(residues 452-486, 487-523, 524-561, and 566-604) and the C-terminal trypsin-fold S1 serine
protease domain (SPD; residues 614-855) with the catalytic triad composed of the residues
Ser805, Asp711, and His656 [47]. Serine proteases of the S1 fold including matriptase display a
pronounced specificity for the positively charged amino acids arginine and lysine at the P1
position [39;46;48-50].
16

Fig 2. Schematic presentation of matriptase.
Matriptase consists of a transmembrane signal anchor domain, a SEA domain; two CUB domains;
4 LDLR domains; and a class S1 serine protease domain.
The extended substrate specificity is determined by the topology of the binding pocket and the
extended specificity profile of matriptase was determined with the solution of the crystal
structure of the serine protease domain of matriptase and by using positional scanning synthetic
combinatorial library (PS-SCL) and substrate phage library. By these methods, the preferred
cleavage sequences (P 4 -P 3 -P 2 -P 1 P 1´) of matriptase was identified to be R/K-XSR A and X-R/K-
SR A, where X is a non-basic amino acid [50-52]. Thus unlike trypsin, matriptase does not
indiscriminately cleave peptide substrates after Lys or Arg but requires recognition of additional
residues. The extended substrate profile correlates very well with the activation motif of
matriptase itself (RQAR VVGG) suggesting that matriptase can be activated by a transactivating
mechanism.
In vitro and cell culture studies have identified several possible substrates of matriptase; prohepatocyte
growth factor/scatter factor (HGF/SF) [53;54], pro-macrophage-stimulating protein 1
(MSP-1) [55], pro-urokinase-type plasminogen activator (uPA) [51;53;56], protease activated
receptor 2 (PAR-2) [51;57], matrix metalloproteinases 3 (MMP-3) [58], pro-kallikreins [59],
immunization M(+)HEp3 associated 135 kDa protein/CUB domain containing protein 1
(SIMA135/CDCP1) [60], insulin-like growth factor binding protein-related protein 1 (IGFBP-rP1)
[61], vascular endothelial growth factor receptor 2 (VEGFR-2) [62], platelet-derived growth factor
(PDGF) [63;64], epidermal growth factor receptor (EGFR) [65;66], fibronectin [26], laminin [26;67]
and gelatine [39].
The function of the intracellular domain is unclear, though data suggest a function in interaction
with the cytoskeleton to regulate localization of matriptase on the cell surface to micro domains
of the plasma membrane through interaction with the actin-associated protein filamin [30;68].
The extracellular non-catalytic domains are likely to function in protein-protein interactions that
modulate localization, activation and inhibition and possibly substrate specificity [28;29;35]. E.g.
in the complement protease cascade, CUB domains are important for the formation of C1r/C1s
tetramers prior to binding of C1q and activation of the C1r and C1s proteases within the complex
[69].
17

Regulation of matriptase
Under physiological conditions proteases are strictly regulated in time and space, thereby
restricting their activity to specific subcellular sites and limiting their access to substrates. This
regulation is achieved by controlled zymogen activation and interaction with inhibitors [2].
Matriptase inhibitors
The serpin family is the largest family of serine protease inhibitors, and complexes between
matriptase and the serpins anti-thrombin III, α1-antitrypsin, and α2-antiplasmin have recently
been purified from human milk and epithelial cell lines, and protease nexin 1 (PN1) complexes
with matriptase was reported in vitro [70-74]. However, the physiological relevance of serpinmediated
inhibition of matriptase has not yet been established. Instead, two members of the
family of Kunitz-type serine protease inhibitors have been identified as essential inhibitors in
regulation of matriptase.
The Kunitz-type inhibitors are a class of serine protease inhibitors that make reversible
interactions with their target proteases [31;75;76]. They are members of the type I
transmembrane protein family and characterized by having one or two extracellular Kunitz-type
serine protease inhibitor domains (KD), a single transmembrane spanning domain and a short C-
terminal cytoplasmic domain [75;76]. They have an N-terminal cleavable signal sequence and a
stop-transfer sequence that halts further translocation through the ER membrane and acts as a
transmembrane anchor [77]. The extracellular part of the membrane bound protein can be
released from the plasma membrane by ectodomain shedding (supplementary I [27]; [75]. While,
HAI-1 has been purified in a complex with matriptase from human milk, seminal fluid and urine
[44;78], complex formation between HAI-2 and matriptase has so far only been observed in vitro
[79].
HAI-1 and HAI-2 were originally identified as potent inhibitors of hepatocyte growth factor
activator (HGFA) from the conditioned medium of the human stomach carcinoma cell line,
MKN45 [80;81].
HAI-1 is the most well studied inhibitor of matriptase and is ubiquitous expressed in epithelia and
predominately localizes to the basolateral surface of epithelial cells, especially the simple
columnar epithelium of ducts, tubules and mucosal surfaces [10;24]. HAI-1 is also expressed in the
endothelial cells of capillaries, venules and lymph vessels [82].
Tissue distribution of HAI-2 has been assessed in both human and mouse. In situ hybridisation of
adult human samples showed a distribution of the inhibitor in epithelial layers of e.g. placenta,
pancreas, kidney and prostate [81]. Tissue distribution of HAI-2 in mice was determined using
enzymatic gene trapping and -galactosidase as a reporter system, and showed a co-localization
of HAI-2 with HAI-1 and matriptase in most epithelial tissue [79]. Additionally, HAI-2 and HAI-2 are
also co-expressed in developing neural tube [79;83].
Human HAI-1 is composed of 513 amino acids with a 35 amino acid signal peptide, thus, the
mature full-length form is composed of 478 amino acids. Full-length HAI-1 has a calculated
molecular weight of 53,319 Da [29;84] and has three potential N-glycosylation sites at Asn66,
Asn235 and Asn507 [80]. HAI-1 is composed of two extracellular Kunitz domains, KD1 (residues
250-300) and KD2 (residues 375-425), an extracellular LDLR domain (residues 319-353), a motif at
18

the N-terminal containing seven cysteines (MANSC) (residues 57-147), a transmembrane domain
(residues 450-472) and a short C-terminal cytoplasmic domain as outlined in fig. 3 [80].
Fig. 3. Schematic presentation of HAI-1.
Domain structure of HAI-1 consists of a MANSC domain, two Kunitz-type domains separated by a
LDLR domain and a transmembrane domain. HAI-1 is a type I membrane-anchored protein.
The overall topology of HAI-2 is similar to that of HAI-1. Human HAI-2 is composed of 252 amino
acids with a 27 amino acid signal peptide, thus, the mature full-length form is composed of 225
amino acids with a calculated molecular mass of 25,415 Da [81]. HAI-2 is composed of two
extracellular Kunitz domains, KD1 (residues 38-88) and KD2 (residues 133-183), and a short C-
terminal region (24 aa) as outlined in fig. 4. HAI-2 has two putative N-glycosylation sites at Asn57
and Asn94 [81].
Fig. 4. Schematic presentation of HAI-2.
Domain structure of HAI-2 consists of a transmembrane domain and two Kunitz-type domains.
HAI-2 is a type I membrane-anchored protein.
The amino acid in the P1 position of the reactive site is of essential importance for the inhibitory
activity of KD [85]. For HAI-1 the essential amino acids in the P1 positions are arginine and lysine
for KD1 and KD2, respectively, and for HAI-2 arginine is present in P1 of both KD1 and KD2
[86;87]. Site-directed mutagenesis studies suggest that KD1 of both HAI-1 and HAI-2 is responsible
for the inhibitory activity against proteases, and for HAI-1 this is also shown for matriptase
[84;87;88].
19

Matriptase proteolytic processing and activation
Matriptase is synthesized as an inactive single-chain zymogen and its activation requires two
sequential proteolytic processing events. The pro-enzyme is first processed at the N-termini of the
stem domain after Gly149 within a conserved G SVIA motif of the SEA domain. This initial
cleavage has been proposed to occur by non-enzymatic hydrolysis of the peptide bond, as it has
been observed for other SEA containing proteins [89]. The second and actual activation site
cleavage event occurs after Arg614 within the highly conserved R VVGG activation motif and is
dependent on the first cleavage at Gly149 [28]. Hereby the disulfide-linked proteolytic active
matriptase is formed (fig. 5). In spite of these processing events, the catalytic domain still remains
attached to the plasma membrane due to the disulfide bond linking it to the stem domain, and
strong hydrophobic interactions within the SEA domain [28;39;51;90]. It is the activity of the
mature enzyme that is believed to be responsible for both the physiological and pathological
functions of the enzyme identified by studies of animal models and human genetics.
For the greater part of serine proteases the activation cleavage depends on another upstream
active protease(s), however evidence indicate that matriptase undergoes autoactivation both in
solution and in the membrane-bound form caused by weak inherent activity of the matriptase
zymogen [28;29;40;91]. First off, the resemblance of the preferred cleavage sequences to the
activation site motif of matriptase [39;50]. Second and most importantly, mutations of any of the
amino acids in the catalytic triad of matriptase (Ser805, Asp711, or His656) result in matriptase
mutants that are processed at Gly149, but unable to undergo the final proteolytic processing at
Arg614. This led to a proposed transactivation mechanism for activation of matriptase [28].
Autoactivation, by which two or more zymogen proteases interact and activate one another by
virtue of their weak intrinsic proteolytic activity, is believed to be relevant for proteases at the
apex of a protease cascades as the active site triad of serine proteases is preformed in the
zymogen protease [92-94]. A well studied example of this mechanism is the activation of
complement C1r protease zymogen where complex formation induce intramolecular
conformational changes that result in activation by a transactivation mechanism [94].
Autoactivation has also been proposed as a mechanism of activation for several other members
of the TTSP family including matriptase-2 and hepsin [45;93].
The activation of matriptase is extraordinary complex, and far from fully understood. Evidence
suggests that the transactivation mechanism for matriptase rely on additional parameters for
matriptase activation to occur including its own multi-domain structure, the plasma membrane as
well as HAI-1. Membrane anchorage of full-length matriptase is required but insufficient for
activation as shown in studies of matriptase in simple vesicle structure implying that a higher
degree of organization within the plasma membrane is indispensable for matriptase activation
[29]. A series of deletion -and point mutations demonstrated that activation of matriptase
requires glycosylation of the first CUB domain (Asn302) and of the catalytic domain, as well as the
presence of intact LDLR domains
[28;29;95]. Thus, autoactivation of matriptase may occur by interaction of two or more
neighboring SEA domain processed matriptase zymogen molecules, and possibly other cofactors
to induce the necessary conformational changes in the substrate binding pocket required for
catalysis [28].
20

Fig. 5. Proteolytic processing of matriptase and inhibition by HAI-1.
Matriptase is synthesized as a 95 kDa Full-length protease that undergoes two sequential
cleavages to become fully active. The first cleavage is after Gly149 in the SEA domain and the
second cleavage is after Arg614 in the conserved activation site motif N-terminal to the serine
protease domain. This processing results in a disulfide linked active form of matriptase. Following
activation, matriptase forms a SDS-resident complex with HAI-1.
After activation, matriptase is inhibited by HAI-1 within a short time rendering active matriptase
little time to act on substrates, see fig. 5 [21;30;31]. Even so, cell culture studies indicate that HAI-
1 also has a role for proper expression, activation and trafficking of matriptase [28;96]. However,
the requirement for HAI-1 in these processes is poorly understood. In breast cancer cells that
express neither matriptase nor HAI-1 endogenously, matriptase is retained in the ER/Golgi
compartments unless the cells are simultaneously transfected with HAI-1. As no difficulties in
trafficking of enzymatically dead matriptase (S805A) was observed in the absence of HAI-1, the
inability of wild-type matriptase trafficking without HAI-1 was suggested to be caused by the
potentially toxic effect of its own unopposed proteolytic activity [96]. Also matriptase activation
was impaired when matriptase was co-expressed with HAI-1 mutated in its LDLR domain [28;96].
Thus, HAI-1 appears at all times to be available for matriptase, which could serve to protect the
cell against aberrant matriptase proteolysis.
The mechanisms triggering matriptase activation remains largely unclear, although data suggest a
role for a number of extracellular stimuli. The signaling sphingolipid sphingosine 1-phosphate
21

(S1P) was identified as a serum component capable of inducing matriptase activation in
immortalized breast epithelial cells and the steroid sex hormone androgen is able to induce a
robust but more slow activation of matriptase in human prostate cancer cells [30;31;97]. In spite
of the divergence in the nature of matriptase activation in different cell types, the subsequent
complex formation with HAI-1 seems to be uniform for both cell types. This is supported by the
identification of suramin as a universal inducer of matriptase activation [30]. Also, exposure to a
mildly acidic extracellular milieu or lowering of ionic strength induces robust and rapid matriptase
zymogen activation in both cell free settings and in several epithelial cell types [29;31;95;98;99].
Although matriptase remains membrane-associated after proteolytic processing and activation it
is clear that matriptase is shed from the plasma membrane given that the protease has been
purified from human milk [44]. N-terminal sequencing of the matriptase isoforms isolated from
conditioned media of an epithelial cell line showed proteolytic cleavage either after Lys189 in the
SEA domain or after Lys204 in the linker region between the SEA domain and the CUB1 domain
[31]. The mechanism(s) by which matriptase is released from the plasma membrane and the
proteases mediating the release remain to be identified. However, cell culture studies suggest
that shedding of matriptase is dependent on the protease being fully proteolytic processed and
possibly complex-formation with HAI-1 [31;78;100].
Prostasin
Prostasin is a GPI-anchored serine protease that was first isolated from human seminal fluid, but
displays a widespread tissue distribution in both mice and humans and is co-expressed with
matriptase in most murine epithelial tissues [9;101;102]. Prostasin is also termed channel
activating protease 1 (CAP-1), following its ability to activate epithelial sodium channel (ENaC)
[103;104]. Full-length human prostasin consists of 343 aa with a 32 aa N-terminal signal peptide
that is proteolytically removed in the ER. Instead prostasin is modified with a C-terminal GPIanchor,
this result in a 40 kDa mature protein that consists of an extracellular serine protease
domain tethered to the outer leaflet of the plasma membrane [102;105], see fig. 6. The serine
protease domain contains the catalytic triad that is formed by His53, Asp102 and Ser206
[102;106]. Prostasin is synthesized as an inactive zymogen but unlike matriptase, prostasin is
incapable of autoactivation and thus requires the proteolytic activity of a second protease to
process the zymogen into the two-chain disulfide-linked active form [105;107]. In vitro matriptase
activates prostasin by proteolytic cleavage in the amino-terminal pro-peptide region of prostasin
at the Arg44 in human prostasin [16;65]. Like matriptase, prostasin has a preference for poly-basic
substrates which also explain the inability of prostasin to autoactivate as the activation site motif
of human prostasin is PQAR ITGG [107]. Prostasin is inhibited by PN1, HAI-1 and HAI-2 like
matriptase [21;108;109].
Prostasin is shed from the apical plasma membrane. In prostate epithelium this secretion is
depended on C-terminal processing at the conserved residue Arg322, while secretion from kidney
and lung epithelial cells depends on GPI-anchor cleavage by the endogenous GPI-specific
phospholipase D1 (Gpld1) [23;101].
22

Fig. 6. Schematic presentation of prostasin.
Prostasin is posttranslationally modified with a GPI-anchor that tethers the single serine protease
domain of prostasin to the membrane.
Physiological functions of matriptase and its role in pathological processes
Matriptase has been shown to play a role in a number of different proteolytic processes at the cell
surface; however, the mechanisms by which matriptase modulates its effect at the molecular
level is still unclear. The importance of matriptase-dependent proteolysis is derived from
combined knowledge from genetic disorders and transgenic animal models, as well as cell culture
studies. Below I will review physiological and pathological processes in which matriptase plays a
crucial role.
Matriptase is crucial for epidermal integrity
In humans, a few studies have been reported identifying mutations in the matriptase gene all of
which lead to various types of the skin disorder ichthyosis [13-15]. The first documented patients
suffer from the rare skin disease autosomal recessive ichthyosis with hypotrichosis (ARIH)
characterized by congenital ichthytosis associated with frizzy hair. The syndrome is caused by
homozygosity for the missense mutation G827R in the serine protease domain of matriptase. The
G827R mutant form of matriptase has a reduced proteolytic activity in vitro as the mutation
affects access to the active site binding cleft [17;18]. Further information about the disease was
obtained by mouse models of the disorder. Xenografting of matriptase-deficient skin onto adult
athymic mice and the generation of a matriptase hypomorphic mouse model both phenocopied
the microscopic hallmarks of the skin from ARIH patients [17;110]. Biochemical analysis of ST14
deficient murine and human epidermis revealed highly reduced prostasin activation and reduced
processing of the abundant epidermal polyprotein pro-filaggrin into filaggrin [13-17;19;110].
Another example of matriptase´s importance in epidermal integrity is its recently shown
implication in Netherton´s syndrome [59]. This disease is a form of ichthyosis characterized by
premature stratum corneum shedding resulting in direct exposure of the living surface of the
epidermis to the external environment and chronic inflammation [111-113]. The genetic
23

ackground of Netherton´s syndrome is a deficiency for the protease inhibitor LEKTI [113]. In a
mouse model of the syndrome, matriptase initiate a run-away kallikrein proteolytic cascade in the
absence of LEKTI that results in an over accumulation of processed filaggrin [59;114].
Additionally, an increase in matriptase expression has been suggested as a common basis for a
range of different human skin disorders [115].
The matriptase-prostasin proteolytic pathway(s) in the epidermis
Matriptase and prostasin are generally co-expressed in terminally differentiated cells of stratified
epithelia that do not possess any proliferative capacity [9]. Studies of both human tissue samples
and murine models as well as cell culture studies have revealed that both matriptase and
prostasin are important factors in regulating terminal epidermal differentiation and hair follicle
development [4;9;12;17;19;116]. The phenotypes observed in mice deficient of prostasin in the
skin are nearly indistinguishable from matriptase deficient mice [4;19;116]. Both mouse models
display compromised epidermal tight junctions and a generalized disruption of the stratum
corneum architecture. The impaired epidermal barrier function is a result of perturbation of
several processes that normally takes place during terminal epidermal differentiation. They
include lipid matrix formation, formation of the water-impermeable cornified layer, and
desquamation (shedding of corneocytes) as well as hyperproliferation of basal keratinocytes, see
fig. 7. These defects result in a lack of terminal epidermal differentiation and are accompanied
with an impaired skin barrier function resulting fatal dehydration and death within 48-60 hours
postnatal [4;19;116].
On the molecular level, matriptase and prostasin deficiency results in greatly reduced proteolytic
processing of pro-filaggrin [15;17;19;116;117]. Studies show that matriptase act upstream of
prostasin in a matriptase-mediated proteolytic activation cascade most likely by directly activating
the prostasin zymogen, which can be seen in fig. 7 [15-17;21]. However, the specific molecular
mechanism by which matriptase and prostasin facilitates profilaggrin processing is unclear. The
hypothesis is supported by the lack of active prostasin in cultured primary keratinocytes from
patients with loss-of-function mutations in ST14 and in the epidermis of matriptase deficient
mice, whereas active prostasin can be readily detected in wild-type human keratinocytes and in
murine epidermis [4;15;16;21;116]. This correlates with activation of prostasin being suppressed
by matriptase ablation in cultured human keratinocytes and that matriptase is able to activate
prostasin in vitro [16;21]. Moreover, the two proteases display synchronized developmental onset
of expression which correlates with acquisition of epidermal barrier function in mice [16].
Together this suggests that in the skin matriptase and prostasin are functioning in the same
pathway and that matriptase act upstream of prostasin in this matriptase-prostasin proteolytic
cascade essential for terminal epidermal differentiation. Furthermore, it has been suggested that
HAI-1 plays an important role in regulating this proteolytic cascade as inhibitor complexes of HAI-
1 with both matriptase and prostasin have been detected in human cultured keratinocytes
induced to differentiate into an organotypic culture model of the skin [21;115].
24

Fig. 7. Matriptase´s role in the epidermis
(A) Matriptase expression is confined to the uppermost living layers of the interfollicular epidermis;
the transitional layer (blue color, arrowhead). This is visualized with a knock-in mouse with a
promoterless β-galactosidase marker gene inserted into the endogenous matriptase gene. The
basal layer is visualized using a keratin-5 antibody (red color, arrow) [12]. Deficiency of either
matriptase or epidermal prostasin leads to a range of epidermal defects including loss of tight
junctions, lipid extrusion and impaired processing of pro-filaggrin that result in impaired epidermal
barrier function (compare B and C). Figure is from [118].
Matriptase and prostasin also co-localize in a number of epithelia other than the epidermis
suggestive of a role for a matriptase-prostasin proteolytic cascade in other organs as well [9].
However, although we know that matriptase and prostasin are linked, we have very restricted
knowledge of the pathway(s) in which they function. Identification of components acting
upstream or downstream of matriptase and prostasin would greatly add to our understanding of
the role(s) of these proteases in vivo.
Matriptase has also been proposed to be at the apex of a matriptase-prostasin proteolytic
cascade activating ENaC that is important for maintenance of salt and water homeostasis by
reabsorption of Na + -ions [119;120]. This channel is also important for normal terminal
differentiation of the epidermis [121]. Both matriptase and prostasin are able to activate ENaC in
Xenopus oocytes [103;120]. So far evidence for a physiological role in channel activation is limited
to prostasin, where fluid clearance from the lungs are impaired in mice with alveolar epitheliumspecific
ablation of prostasin [122]. Intriguingly, the catalytic activity of prostasin is dispensable
for its capability to activate ENaC in the Xenopus model system. Catalytically inactive prostasin
induce both ENaC cleavage and activation, nonetheless cell surface expression of prostasin is still
essential for activity [123;124].
Matriptase has an important role in epithelial homeostasis
The generation of ST14 (the gene encoding matriptase) deficient mouse models established that
matriptase has a critical role in the development and maintenance of multiple epithelial tissues
25

consistent with its widespread epithelial distribution [4;5;12]. Embryonic tissue specific ablation
or acute ablation of matriptase in adult tissues of mice results in severe organ dysfunction and
widespread epithelial demise, often associated with increased paracellular permeability, loss of
tight junction function and mislocation of tight junction-associated proteins, see fig. 8. Ultimately
these defects cause a reduction in body weight and ultimately death [5]. Matriptase deficiency in
mice also affects hair follicle development and results in dramatically increased thymocyte
apoptosis, and depletion of thymocytes [4;12].
Fig. 8. Matriptase´s role in the intestinal epithelia
(A) Matriptase is expressed in goblet cells (arrow) and in surface mucosal cells (arrowhead) in the
murine intestine. Sustained matriptase expression is essential for maintenance of the intestinal
epithelia. Ablation of matriptase disrupts tight junctions and cell polarity (compare B to C) and
causes architectural distortion and compromised barrier function in the large intestine resulting in
edema and diarrhea leading to premature death in mice. Figure is from [118].
Matriptase is proposed to promote intestinal barrier recovery in injured intestinal mucosa of
inflammatory bowel disease (IBD). Studies of human specimens and mouse models show that
matriptase is down-regulated in inflamed colonic tissues from IBD patients and that matriptase
has a protective role against colitis in mice [8;125]. Thus, matriptase has an essential role in
development as well as maintenance of multiple types of epithelia.
Importance of matriptase regulation in vivo
Under normal physiological conditions, proteases are strictly regulated at the protein level.
Synthesis of proteases as inactive zymogens and complex formation with protease inhibitors limit
their accessibility to substrates. The importance of protease inhibitors to govern protease activity
is obvious, as the activation zymogens to active proteases is an irreversible action. Therefore,
important knowledge of proteases can be obtained indirectly by knock-out studies of their
respective inhibitors. This section presents important knowledge about matriptase obtained by
studies of knock-out animal models of its physiological inhibitors, HAI-1 and HAI-2.
26

Even though matriptase is widely expressed in both embryonic and adult epithelial tissues the
protease does not have critical non-redundant functions until after birth [4]. Surprisingly, the
proteolytic activity of matriptase must nevertheless be under strict control during embryogenesis
for correct placental development to take place. Ablation of HAI-1, HAI-2, or the combined
haploinsufficiency for HAI-1 and HAI-2 leads to failure of placental labyrinth formation [15;126-
128]. Correct placental development can be restored with simultaneous ablation of matriptase,
indicating that the placental defects in HAI-1-deficient, HAI-2-deficient or HAI-1/HAI-2
haploinsufficient mice are caused solely by unopposed matriptase activity [83;126-128]. Thus
although matriptase is completely dispensable for embryogenesis, its activity needs to be strictly
regulated by HAI-1 and HAI-2 for placental development to take place.
HAI-1-mediated inhibition of matriptase also has an essential role in postnatal epithelial
homeostasis. The generation of a chimeric mouse model with sufficient placental HAI-1, but
ablation of HAI-1 in the embryo resulted in viable offspring [20;129]. These mice suffer from
various defects of the keratinized epithelium and die before adulthood. The same genotype
superimposed on hypomorphic expression of matriptase completely eliminates epidermal barrier
and hair follicle defects in HAI-1 deficient mice [20;129]. Similar outcome is reported for
zebrafish, where deletions of the HAI-1 orthologues Hai-1a and Hai-1b result in skin inflammation
and a compromised epithelial integrity caused by detrimental matriptase activity that result in
death app. 24 hrs after fertilization [22].
Excess local matriptase activity is detrimental not only to mouse placental labyrinth formation but
also for neural tube closure, which became evident with the generation of mice deficient for HAI-
2. HAI-2 and matriptase are expressed in the neural and non-neural ectoderm precisely at the
interface where the two neural folds fuse. Simultaneously ablation of matriptase however, only
partially rescues this HAI-2 knock-out phenotype suggestive of several in vivo target of HAI-2 in
neural tube closure [83]. In humans, a range of mutations in the spint2 gene encoding HAI-2 have
been reported. These patients present a wide range of developmental abnormalities including
duplication of internal organs and digits, craniofacial dysmorphisms, as well as anal and choanal
atresia [130]. It remains to be established if these defects are caused by excess matriptase activity
or unregulated activity of other serine proteases.
Hence, strict and proper regulation of matriptase is important for postnatal survival, development
and maintenance of several epithelia and partially for neural tube closure as these defects were
caused by unregulated matriptase activity.
Matriptase in carcinogenesis
85 % of all cancers originate from epithelial cells that line the internal and external surfaces of the
body. During transformation of normal epithelia into cancerous tissue a number of events occur;
function-altering mutations of oncogenes and tumor suppressor genes, loss of epithelial cell
polarity, concomitant tissue disorganization, penetration of the basement membrane and
invasion of transformed epithelial cells into the underlying stroma [131]. Unlike most proteases
involved in carcinogenesis which are expressed by the connective tissue supporting the epithelia,
matriptase is expressed by the transformed epithelial cells themselves [6]. Matriptase is
undoubtedly involved in carcinogenesis, however at present its role remains obscure. Knowledge
27

of matriptase´s role in cancer comes from studies of animal models, tissue samples and cancer
cell lines.
The oncogenic potential of matriptase was directly shown in transgenic mice, where a modest
over-expression of wild-type matriptase in the skin caused spontaneous squamous cell carcinoma
formation to occur. Moreover, matriptase supports both ras-dependent and independent
carcinogenesis and potentiate the effects of genotoxic exposure [6]. Matriptase expression
undergoes a dramatically spatial redistribution during the transition of epidermal lesions from
hyperplasia to dysplasia and becomes present in the proliferating basal compartment. Hereby
matriptase is able to mediate c-Met induced activation of the PI3K-Akt-mTor pathway through
matriptase-catalysed HGF activation and binding of the HGF ligand to the c-Met receptor [6;7;12].
Simultaneously ablation of cMet or a corresponding over-expression of matriptase´s cognate
inhibitor HAI-1 negated the oncogene potential of matriptase over-expression indicating the
impact of unopposed protease activity in malignant transformation [6;7]. In addition, matriptase
has been shown in vitro to activate several molecules associated with cancer progression
including pro-HGF/SF [53], pro-MSP-1 [55], pro-uPA [51;53;56], PAR-2 [51;57], MMP-3 [58],
SIMA135/CDCP1 [60], IGFBP-rP1 [61], VEGFR-2 [62], PDGF [63;64] and EGFR [65;66].
Conversely, a recent study assigns matriptase with a role as tumor suppressor in a murine model
of colitis-associated colon cancer. Intestinal-specific epithelial ablation of matriptase resulted in
intrinsic intestinal permeability barrier perturbations that progressed into chronic inflammation
and subsequently formation of colon adenocarcinoma highlighting the importance of matriptase
in epithelial tight junction formation [8].
Multiple studies have also assessed the mRNA and protein levels of matriptase during human
carcinogenesis and the potential value of matriptase as a novel prognostic marker, a predictor of
patient outcome and as a possible therapeutic target for human cancers. However, the findings
from these studies are ambiguous.
For some cancers, an increase in matriptase mRNA or protein expression levels has been
associated with tumor progression in e.g. breast, cervical, ovarian and prostate cancer [132-139].
In contrast, other studies report a significant downregulation of matriptase expression levels in
gastric, colorectal and breast cancer [140-142]. In fact, one of the groups first to discover
matriptase did so by identifying genes down-regulated in human colorectal carcinomas, hence
matriptase gene annotation; suppressor of tumorigenicity-14 [41;143].
Explanation for these discrepancies could be explained by different roles of matriptase in different
epithelia and cancer types and partly by differences in tissue sampling, tissue composition, tumor
staging and differences in quantification methodology. Expression studies including both
matriptase and HAI-1 may be more informative as HAI-1 has important roles in matriptasemediated
carcinogenesis in mice and for epithelial integrity in general [6;22;128;144]. An
imbalance in the matriptase-HAI-1 ratio resulting in a larger proportion of free active matriptase
could lead to harmful overactivation of matriptase-mediated pathways contributing to
carcinogenesis [145]. This proposal of an imbalance in the matiptase-HAI-1 ratio is supported by
studies in gastrointestinal, colorectal, and ovarian cancers where the protease-inhibitor ratio was
shifted in favor of matriptase when comparing advanced stage tumors with low-stage lesions or
corresponding tissue from control individuals [140;141;145;146].
28

Technical considerations
This chapter includes methodological considerations for the main methods used for the work
presented in three manuscripts; paper I, II and III.
Cell system
Matriptase, prostasin and HAI-1 are all expressed in polarized epithelia, yet matriptase and HAI-1
are located to the basolateral plasma membrane at steady state and prostasin locates to the
apical plasma membrane at steady state [9;11;24;102]. However, the organization of the
epithelium makes it difficult to study the polarity of the cells and the intracellular trafficking of
proteins since the basolateral plasma membrane cannot be accessed. To obtain this kind of
information it is applicable to utilize model systems of the polarized epithelia. Epithelial cell lines
have proven to be useful tools for studying epithelium.
We have previously used the Mardin Darby canine kidney (MDCK) cell line as a model system for
polarized epithelia to delineate the intracellular trafficking of HAI-1 (supplementary I; [27]). In the
present study (paper I and III), we decided to use the well known Caco-2 cell line as a model
system for the polarized epithelium. MDCK and Caco-2 cells have both been extensively used as
models of the polarized epithelium [147;148]. However combined with the available antibodies,
the Caco-2 cell line offers the advantage that we can examine endogenously expressed
matriptase, prostasin and HAI-1, which eliminates unwanted effects by recombinant
overexpression.
Caco-2 cells originate from a human colon carcinoma, yet in culture these cells spontaneously
differentiate and form a polarized monolayer of cells that morphologically and functionally
resemble the mature enterocytes of the small intestine. For this reason, the Caco-2 cell line has
been extensively used as a model of the polarized epithelium [147]. When cultured onto
Transwell filters, the apical and basolateral plasma membrane, divided by formation of tight
junctions, can be accessed separately, which allows for investigations of intracellular trafficking of
proteins (fig. 9).
Thus, Caco-2 cells grown on Transwell filters are a good model system for studying the
intracellular transport of matriptase, prostasin and their common inhibitor HAI-1. In our studies
(paper I and III) we have used 11 days post-confluent Caco-2 cells grown on Transwell filters as
the level of HAI-1 complexed matriptase reaches a plateau at this differentiation level
(unpublished results, Stine Friis).
Tight junction formation and barrier function of Transwell filter grown Caco-2 cells can be
assessed in several ways. It can be measured by transepithelial electrical resistance (TEER) also;
tracer molecules can be used to measure the permeability of the Caco-2 monolayer, e.g. lucifer
yellow or phenol red [149;150]. However, a much simpler way of evaluating barrier function of
cells grown on Transwell filters is to assess whether the monolayer of cells is able to maintain a
difference in medium level between the inner and outer chamber over night [27].
29

Fig. 9. Monolayers of polarized Caco-2 cells on porous filters.
Caco-2 cells spontaneously form a polarized monolayer of cells when grown on Transwell filters. At
confluence, the cells form tight junctions that separate the apical and basolateral extracellular
spaces. This setting allows for separate access to the apical and basolateral plasma membrane
domains and media. The figure shows a cross section of the Transwell filter insert in the culture
dish.
Although cell lines are extremely useful for obtaining new knowledge about molecular mechanism
responsible for different phenomena, data from one cell line is not always comparable with data
from other cell line nor can one expect data to be transferable to whole organism settings. One
reason for this is the immortalized feature of most cell lines. However, cell lines make good model
systems to study important biological pathways.
Protease pull down assays
Variations over protease inhibitor mediated pull down were applied in paper I and II. These
techniques were used to identify activation state and to assay for the presence of active protease.
Different set ups were used in the two studies but they rely on the same concept.
Co-immunoprecipitation (co-IP) is a well established method for small scale purification of
antigens and their interaction partners and allows for identification of activation status of
proteins. In a co-IP, the target antigen has the function of bait and is used to co-precipitate a
binding partner/protein complex (prey) from a crude cell lysate or tissue extract. An antibody
against the bait is incubated with a lysate either as pre-immobilized onto an insoluble support or
as a free un-bound antibody in combination with an insoluble support e.g. protein G sepharose as
outlined in fig. 10. The main disadvantage of free antibody protocol for immunoprecipitation and
co-IP is that the conditions used to elute the precipitated antigen also release the antibody.
Depending on the size of the antigen, the heavy and light chains of the antibody can completely
30

mask the detection of the antigen/interaction partners in Western blot analysis. However this is
circumvented by cross-linking the antibody to the resin.
Fig. 10. Immunoprecipitation
A suitable antibody is added to a cell lysate or a tissue extract either pre-immobilized onto an
insoluble support or as a free unbound antibody to bind the protein of interest. Gentle incubation
allows the antigen to by immobilized and purified by means of the antibody and the insoluble
support. In the pre-immobilized antibody approach the antibody has been cross-linked to the
insoluble support. Immunoprecipitation of intact protein complexes is known as co-IP.
Immunoprecipitated proteins and their binding partner(s) can be detected by SDS-PAGE and
Western blot analysis. Figure is from [151].
31

We applied co-IP to determine the presence activated matriptase and activated prostasin in
extracts of murine placentas in paper II, as simple detection of the activated proteases in lysates
of placentas was unsuccessful by direct Western blotting. We used a free antibody protocol (the
latter approach) to co-IP matriptase and prostasin with a HAI-1 antibody and used protein G
sepharose that binds to the Fc region of immunoglobulin G as the insoluble support. As unspecific
binding to the protease G sepharose can occur, the lysates were pre-incubated with protein G
sepharose alone to minimize unspecific binding in the actual co-IP reaction. Placentas from
matriptase deficient and prostasin deficient embryos were used as negative controls, respectively.
Using co-IP it is often anticipated that associated proteins (prey) is related to the function of bait
protein. However, this is merely an assumption that needs further verification. Though, in this
case the relationship between matriptase, prostasin and HAI-1 is already well-established [16;21].
Thus, we can use the technique to identify the presence matriptase and prostasin activation in
placental extracts from embryos of different genotype.
In paper I, we use a derivation of immunoprecipitation to detect the presence of active matriptase
and prostasin. Instead of using antibodies to immobilize matriptase and prostasin, we applied the
general serine protease inhibitors aprotenin and leupeptin assuming that only active serine
proteases and not their zymogen counterparts will be able to bind to these serine protease
inhibitors. As HAI-1 forms reversible inhibitor complexes with proteases, immobilized aprotenin
and leupeptin could potentially compete with HAI-1 to bind matriptase and prostasin. This can be
ruled out by a complicated experimental setting where the inhibitor-coupled pull down is
performed on a sample in which matriptase-HAI-1 complex formation is induced. The major
disadvantage of this method is that detection of active matriptase occurs in solution which may
deviate from live cell culture settings.
Biotinylation assays
The highly specific interaction of biotin with avidin/streptavidin is a useful tool in designing
nonradioactive purification and detection systems. The extraordinary affinity of avidin
/streptavidin for biotin is the strongest known noncovalent interaction of a protein to a ligand and
allows biotin-containing molecules in a complex mixture to be isolated by exploiting this highly
stable interaction. For paper I and III included in this thesis, we have exploited the advances of the
biotin-avidin interaction to study matriptase, prostasin and HAI-1. In paper I, we have made use of
biotinylation assays for studying the subcellular trafficking of matriptase, prostasin and HAI-1 in
Transwell filter grown polarized Caco-2 cells. In paper III, we have exploited the
streptavidin/avidin–biotin interaction to extract proteases with a peptidolytic activity towards a
biotinylated chloromethyl ketone peptide inhibitor designed with a preferred substrate sequence
of matriptase.
32

Fig. 11. Reaction of S-NHS-SS-biotin with primary amine on proteins.
Primary amines of proteins react with NHS-esters by nucleophilic attack, whereby the N-
hydroxysulfosuccinimide (NHS) is released as a byproduct. The NHS ester group on this reagent
reacts with the ε-amine of lysine residues and forms a stable product. Hydrolysis of the NHS-ester
competes with the reaction in aqueous solution and increases with increasing pH. α-amine groups
present on the N-termini of peptides also react with NHS esters but these α-amines are seldom
accessible for conjugation in proteins. Figure is from [152].
Biotin is a relatively small molecule and can thus be conjugated to many proteins without
significant affect on the target proteins biological activities. More biotin molecules can bind to a
single protein which greatly increases the sensitivity of many assay procedures. Additionally,
biotin has been modified in numerous ways to accommodate particular applications. In paper I,
we have utilized the membrane impermeable biotin derivative sulfosuccinimidyl-2-
[biotinamido]ethyl-1,3-dithiopropionate (S-NHS-SS-biotin) because this conjugate is cell
membrane impermeable due to the charged sulfonate group and is thus favorable for cell surface
biotinylation. The thiol-cleavable spacer arm of S-NHS-SS-biotin harbors two important
properties. First, the extended spacer arm reduces steric hindrance associated with
avidin/streptavidin binding and thereby enhances the interaction. Second, the S-S cleavable
spacer arm allow for reversible biotinylation, by treatment with reducing agents, which is a
required feature when performing transport studies of cell surface proteins. Specifically, we use
this feature to determine the endocytosis and transcytosis properties of matriptase, prostain and
HAI-1. Finally, S-NHS-SS-biotin reacts with primary amines readily available on the cell surface of
proteins (fig. 11).
33

Fig. 12. Assay for detection of active matriptase in cell culture
To establish an assay for the detection of active matriptase, we developed a synthetic peptide with
binding preference for the active cleft in matriptase. This peptide is coupled to both a biotin-group
and to a chloromethyl ketone (Cmk) group. When the Cmk group is in close proximity to proteases,
covalent bond formation occurs by alkylation of the active site histidine. Active proteases with
binding preferences for the Cmk inhibitory peptide is labeled and extracted by streptavidin pull
down of the biotin group. Specificity for matriptase is obtained via Western blot analysis and
matriptase specific antibodies.
For paper III, we make use of the N-terminal biotin moiety of the CMK peptide inhibitor, biotin-
RQRR-Cmk, as a tag to extract peptidolytic active matriptase from Caco-2 cell lysates. The peptide
sequence of the tetra peptide was designed to obtain the highest specificity towards matriptase
and was deduced from a preferred substrate sequence of matriptase [50]. This peptide enables us
to biotin-label peptidolytic active matriptase in live and intact Caco-2 cells (fig. 12), as opposed to
the inhibitor-coupled sepharose used in paper I that label active matriptase in solution.
Once biotin is attached to its target molecule, the molecule can be immobilized using biotin
binding molecules. In this study we have made use of streptavidin (paper I and III) and monomeric
avidin (paper I). The high affinity biotin-streptavidin binding allows for extraction of biotin from
very complex mixtures, and dissociation between biotin and streptavidin requires harsh
denaturing conditions e.g. boiling in SDS-PAGE sample buffer. In contrast, monomeric avidin binds
biotin in a reversible manner, and gentle recovery of biotinylated molecules is feasible by elution
with free excess biotin. Monomeric avidin can facilitate purification of functional proteins and in
paper I, we employ this in recovery of the SDS-resistant matriptase-HAI-1 complex in Western blot
analysis.
34

Results
Data included in this PhD thesis is presented in the following three manuscripts:
Manuscript I
Transport via the transcytotic pathway makes prostasin available as substrate for matriptase
Manuscript II
Reduced prostasin (CAP1/PRSS8) activity eliminates HAI-1 and HAI-2 deficiency associated
developmental defects by preventing matriptase activation
Manuscript III
Novel assay for detection of active matriptase
35

Paper I
Transport via the transcytotic pathway makes prostasin available as substrate for matriptase
Stine Friis 1 , Sine Godiksen 2 , Jette Bornholdt 1 , Joanna Selzer‐Plon 1 , Hanne Borger Rasmussen 3 ,
Thomas H. Bugge 4 , Chen‐Yong Lin 5 and Lotte K. Vogel 1
1 Department of Cellular and Molecular Medicine, University of Copenhagen, Copenhagen,
Denmark.
2 Department of Biology, University of Copenhagen, Copenhagen, Denmark.
3 Department of Biomedical Science, University of Copenhagen, Copenhagen, Denmark
4 Proteases and Tissue Remodeling Unit, National Institute of Dental and Craniofacial Research,
National Institutes of
Health, Bethesda, USA
5 Department of Biochemistry and Molecular Biology, Greenebaum Cancer Centre, University of
Maryland, Baltimore,
USA
Published in Journal of Biological Chemistry, February 2011.
36

THE JOURNAL OF BIOLOGICAL CHEMISTRY VOL. 286, NO. 7, pp. 5793–5802, February 18, 2011
Printed in the U.S.A.
Transport via the Transcytotic Pathway Makes Prostasin
Available as a Substrate for Matriptase *
Received for publication, September 29, 2010, and in revised form, December 1, 2010 Published, JBC Papers in Press, December 10, 2010, DOI 10.1074/jbc.M110.186874
Stine Friis ‡ , Sine Godiksen § , Jette Bornholdt ‡ , Joanna Selzer-Plon ‡ , Hanne Borger Rasmussen , Thomas H. Bugge ,
Chen-Yong Lin**, and Lotte K. Vogel ‡1
From the Departments of ‡ Cellular and Molecular Medicine, § Biology, and Biomedical Science, University of Copenhagen,
2200 Copenhagen, Denmark and the Proteases and Tissue Remodeling Unit, NIDCR, National Institutes of Health,
Bethesda, Maryland 20892, and the **Department of Biochemistry and Molecular Biology, Greenebaum Cancer Center,
University of Maryland, Baltimore, Maryland 21201
Thematriptase-prostasinproteolyticcascadeisessentialfor
epidermaltightjunctionformationandterminalepidermal
differentiation.Thisproteolyticpathwaymayalsobeoperative
inavarietyofotherepithelia,asbothmatriptaseandprostasin
areinvolvedintightjunctionformationinepithelialmonolayers.However,inpolarizedepithelialcellsmatriptaseismainly
locatedonthebasolateralplasmamembranewhereasprostasinismainlylocatedontheapicalplasmamembrane.Todeterminehowmatriptaseandprostasininteract,wemappedthe
subcellularitineraryofmatriptaseandprostasininpolarized
colonicepithelialcells.Weshowthatzymogenmatriptaseis
activatedonthebasolateralplasmamembranewhereitisable
tocleaverelevantsubstrates.Afteractivation,matriptase
formsacomplexwiththecognatematriptaseinhibitor,hepatocytegrowthfactoractivatorinhibitor(HAI)-1andisefficientlyendocytosed.Themajorityofprostasinislocatedon
theapicalplasmamembranealbeitaminorfractionofprostasinispresentonthebasolateralplasmamembrane.Basolateral
prostasinisendocytosedandtranscytosedtotheapicalplasma
membranewherealongretentiontimecausesanaccumulationofprostasin.Furthermore,weshowthatprostasinonthe
basolateralmembraneisactivatedbeforeitistranscytosed.
Thisstudyshowsthatmatriptaseandprostasinco-localizefor
abriefperiodoftimeatthebasolateralplasmamembraneafterwhichprostasinistransportedtotheapicalmembraneas
anactiveprotease.Thisstudysuggestsapossibleexplanation
forhowmatriptaseorotherbasolateralserineproteasesactivateprostasinonitswaytoitsapicaldestination.
Thetrypsin-likemembraneserineproteasematriptaseis
essentialformaintenanceofmultipletypesofepithelia.ConditionalablationoftheSt14genecodingformatriptasein
intestine,kidney,andlungofadultmiceresultsinweightloss,
severedeclineinhealthanddeathwithin2weeks,causedby
* This work was supported, in whole or in part, by the NIDCR Intramural
Research Program and by National Institutes of Health Grant R01-CA-
123223. This work was also supported by The Harboe Foundation, The
Augustinus Foundation, The Brothers Hartmanns Foundation, The A.P.
Møllers Foundation for the Advancement of Medical Science, The Cluster
of Cell Biology at the University of Copenhagen, and the Lundbeck
Foundation.
1 To whom correspondence should be addressed: Blegdamsvej 3, Bldg. 6.4,
2200 Copenhagen N, Denmark. Tel.: 45-35-32-77-87; Fax: 45-35-36-79-80;
E-mail: vogel@sund.ku.dk.
organdysfunctionassociatedwithincreasedpermeabilityand
lossoftightjunctions(1).Knockdownofmatriptaseby
siRNAinacellmodeloftheintestinalepitheliumcauseda
leakybarrier,impairedabilitytodeveloptransepithelialelectricalresistance(TEER)
2 andenhancedparacellularpermeabilitythroughregulationoftightjunctionproteins(2).Togetherthesedatasuggestakeyroleformatriptaseinepithelial
barrierfunctionandtightjunctionassembly.
Prostasin(alsoknownasCAP1andPRSS8)isaGPI-anchoredtrypsin-likeserineprotease.Prostasinisco-expressed
withmatriptaseinmostepithelialtissuesincludingtheepidermis,kidney,andcolon(3).Prostasinproteolyticactivity
hasalsobeensuggestedtopromotethedevelopmentoffunctionaltightjunctions,TEERandparacellularpermeability
(4–6).Unlikematriptase,whichundergoesefficientautoactivation,theprostasinzymogenisnotabletoauto-activate,
andformationofactiveprostasinrequiresactivationsite
cleavagebyothertrypsin-likeserineproteases.Strongdata
suggestthatmatriptaseactsupstreamofprostasininazymogencascadeintheepidermis.Thesevereepidermaldefectsof
matriptasedeficiencyappeartobeaconsequenceoflackof
activeprostasin.Onlytheinactiveformofprostasinisfound
inmatriptase-deficientmiceandmatriptase-deficientand
prostasin-deficientmicehavenearlyidenticalphenotypes
withcompromisedepidermaltightjunctionformationandno
terminalepidermaldifferentiation(6–11).Furthermore,it
hasbeenshowninvitrothattheserineproteasedomainof
matriptaseisdirectlyabletocleavethezymogen-formof
prostasin,togenerateproteolyticallyactiveprostasin(11).
Matriptase-dependentactivationofprostasinwasrecently
demonstratedinahumanorganotypicskinmodelandin
matriptase-deficienthumanepidermis(6,12).
Theplasmamembraneofapolarizedepithelialcellisdividedintoanapicalandabasolateralplasmamembranedomainseparatedbytightjunctions.Thetightjunctionsprevent
diffusionofmembraneproteinsbetweenthetwomembrane
domains.Inpolarizedepithelialcellstherearetwopathways
fornewlysynthesizedproteinstoreachtheapicalplasma
membrane:ThedirectpathwayfromthetransGolginetwork
directlytotheapicalplasmamembraneandtheindirectpath-
2 The abbreviations used are: TEER, transepithelial electrical resistance;
HAI-1, hepatocyte growth factor activator inhibitor 1; ENaC, epithelial
sodium channel.
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FEBRUARY 18, 2011 • VOLUME 286 • NUMBER 7 JOURNAL OF BIOLOGICAL CHEMISTRY 5793

Matriptase Activates Prostasin on the Basolateral Plasma Membrane
way,wherenewlysynthesizedproteinsviathebasolateral
plasmamembraneareendocytosedandtranscytosedtothe
apicalplasmamembrane(13).Ithasbeenreportedthat
matriptaseismainlylocatedatthebasolateralplasmamembraneinratenterocytesandotherpolarizedepithelialcells
(14,15).However,proteolyticallyshedmatriptaseincomplex
withHAI-1waspurifiedfromhumanmilksuggestinganapicalsecretion(16).Conversely,thematriptasesubstrate,prostasin,ismainlylocatedattheapicalplasmamembraneofpolarizedepithelialcells(17,18).
Thepresentstudyaimstodeterminewhereinthepolarized
epithelialcellactivematriptaseinteractswithitssubstrate
prostasin,inordertoexplainhowabasolateralproteasecan
cleaveandactivateanapicallylocatedsubstrate.Matriptaseis
synthesizedasaninactive,single-chainzymogen.Itsactivationrequirestwosequentialendoproteolyticcleavages.The
firstproteolyticprocessingcleavageoccursafterGly-149,yet
theprocessedformremainstightlyassociatedwiththe
membrane.
Matriptaseis,subsequently(anddependentonthefirst
cleavage)cleavedafterArg-614intheserineproteasedomain
togainproteolyticactivity.Shortlyafteractivation,matriptase
formsacomplexwithHAI-1,wherebymatriptaseisenzymaticallyinhibited.Hence,substratesshouldbepresentatthe
samelocationasmatriptaseactivationtakesplace.
Wepresentbiochemicaldatashowingthatamatriptaseprostasinzymogencascadeisindeedpossibleinpolarized
epithelialcells.Weshowthatmatriptaseiscleavedtoitsactiveformonthebasolateralplasmamembrane,subsequently
inhibitedbyHAI-1followedbyendocytosis.Importantly,we
findprostasinpresentonthebasolateralplasmamembrane
duringmatriptaseactivation.Furthermore,thebasolateral
prostasinisactiveandtranscytosedtotheapicalplasma
membranewhereitaccumulates.Theseresultsdemonstrates
thatmatriptaseandprostasinmayfunctionallyinteractin
polarizedepithelialcells,despitetheir,respective,basolateral
andapicallocationsatsteadystate.
EXPERIMENTAL PROCEDURES
CellCulture—Caco-2cellsweregrowninminimalessential
mediumsupplementedwith2mM L-glutamine,10%fetalbovineserum,1nonessentialaminoacids,100units/mlpenicillin,and100
g/mlstreptomycin(Invitrogen)at37°Cinan
atmosphereof5%CO 2 .Forallexperiments,210 6 cells
wereseededinto0.4 m-pore-size24mmTranswellfilter
chamber(Corning)allowingseparateaccesstotheapicaland
basolateralplasmamembrane.Cellsweregrownuntilday11
postconfluencebeforetheywereusedforexperiments.The
tightnessoffilter-growncellswasassayedbyfillingtheinner
chambertothebrimandallowingittoequilibrateovernight.
Thecellculturemediumwaschangedeveryday.
Biotinylation,Internalization,andBiotin-removal—Caco-2
cellsgrownontranswellfilterswerewashedthreetimeswith
ice-coldPBS(PBSsupplementedwith0.7mMCaCl 2 and
0.25mMMgCl 2 )onbothapicalandbasolateralside.Thecells
werebiotin-labeledfor30minat4°C,eitherfromtheapical
orthebasolateralside,with1mg/mlEZ-link TM Sulfo-NHS-
SS-Biotin(Pierce)dissolvedinPBS.Afterbiotinlabelingthe
cellswerewashedtwicewithice-coldPBS.Residualbiotin
wasquenchedfor5minat4°Cwith50mMglycineinPBS
andthecellswerewashedagainwithPBS.Forinternalizationexperiments,preheatedmedia(serum-freeMEMmediumcontaining20mMNaHCO
3 ,2mM L-glutamine,100
units/mlpenicillin,and100 g/mlstreptomycin(Invitrogen))
wasaddedandthecellswereincubatedat37°Ctoregainnormaltrafficking.Forendocytosisandtranscytosisexperiments,
surfacebiotinwasremovedafterincubationasindicatedusingthenon-membranepermeablereducingagentglutathione
(SigmaAldrich).Surfaceexposedbiotinwasremovedfrom
eitherapicalorbasolateralsidewith16mg/mlglutathionein
75mMNaCl,75mMNaOH,1mMEDTA,and0.5%BSAin
H 2 Oundergentleagitationfor220min.Cellswerewashed
withPBSandresidualglutathionewasinactivatedwith5
mg/mliodoacetamide(SigmaAldrich)inPBSfor5min.A
setofparallelsampleswasleftwithouttheglutathionereductiontomonitorthetotalamountofbiotinylatedproteinpresentthroughthecourseoftheexperiment.Cellswerewashed
twiceinPBS,andlysedinPBScontaining1%TritonX-100,
0.5%deoxycholateandproteaseinhibitors(10mg/literbenzamidine,2mg/literpepstatinA,2mg/literleupeptin,2mg/l
antipain,and2mg/literchymostatin).Forinhibitor-Sepharosepull-downs,proteaseinhibitorswereomittedfromthe
lysisbuffer.
MonomericAvidin/StreptavidinPrecipitationofBiotinylatedProteins—Lysatesfrombiotin-labeledcellswerecentrifugedat20,000
gfor20mintopellettheinsolublematerial.
Thesupernatantwastransferredtocleaneppendorftubes
witheitherPiercemonomericavidinagarose(Pierce)(120
l/24mmfilter)orPiercestreptavidinagarose(50 l/24
mmfilter),preparedasdescribedbymanufacturer.After
overnightincubationat4°Cwithend-over-endrotation,the
agarosewaswashedthreetimeswith50mMTris-HCl,pH
6.25.Biotinylatedproteinswereelutedfromthemonomeric
avidinagarosewith4mMbiotin(Pierce)inPBSfor30min
followedbyadditionofSDSsamplebuffer.Forelutionfrom
streptavidin-agarose,thesampleswereboiledinSDSsample
buffer.
WesternBlot—The2SDSsamplebuffer(125mMTris-
HCl,25mMEDTA,pH6.8,4%SDS,5%glycerol,0.01%bromphenolblue)didnotcontainanyreducingagentandthesampleswerenotboiledpriortoSDS-PAGEtopreventprotein
complexesfromdissociating,unlessotherwisespecified(0.1 M
dithiothreitol,DTT).AlllysateswereincubatedwiththeSDS
samplebufferfor10minatroomtemperaturebeforegel
loading.Theproteinswereseparatedon7%acrylamidegels
madeinthelaboratoryandtransferredtoImmobilon-PPVDF
membranes(Millipore).Themembraneswereblockedwith
10%nonfatdrymilkinPBScontaining0.1%Tween-20
(PBST)for1hatroomtemperature.TheindividualPVDF
membraneswereprobedwithprimaryantibodiesdilutedin
1%nonfatdrymilkinPBSTat4°Covernight.Thenextday
themembraneswerewashed3withPBSTandthebinding
ofprimaryantibodieswasfollowedbyrecognitionwithsecondaryhorseradishperoxidase(HRP)-conjugatedsecondary
antibodies(Pierce).After3washwithPBST,thesignalwas
developedusingtheECLreagentSuperSignalWestFemto
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Matriptase Activates Prostasin on the Basolateral Plasma Membrane
MaximumSensitivitySubstrate(Pierce),accordingtotheprotocolsuppliedbythemanufacturerandvisualizedwithaFuji
LAS1000-camera(Fujifilm,SwedenAB).Forgraphs,thefree
onlinesoftwareImageJ(createdbyWayneRasband,NIH,
Bethesda)wasusedtoquantifythebandsontheblot.Values
inthegraphrepresentthesumofallthebandsvisualizedon
thegel.Furthermore,thegraphrepresentsthemeanofthree
independentexperimentsandispresentedwiththestandard
deviation.
Antibodies—Theantibodiesusedweremonoclonalmouse
anti-humanantibodiesM32andM69(19).TheantibodyM32
detectsallformsofmatriptase,includingzymogen,activated
form,andcomplexes.Undertheconditionsusedinthisstudy,
theantibodyM69onlydetectedthematriptase-HAI-1complexandM69reactivematerialishereafterreferredtoasthe
matriptase-HAI-1complex.Theotherantibodiesusedwere
polyclonalrabbitanti-humanmatriptaseraisedagainstthe
serineproteasedomainofmatriptase(Cat.no.IM1014,Calbiochem),mouseanti-humanHAI-1antibodyM19(19),and
mouseanti-humanprostasinantibody(Cat.no.612173,BD
TransductionLaboratories).M32,M69,andpolyclonalgoat
anti-humanHAI-1(cat.no.AF1048,R&D)antibodieswere
usedforimmunocytochemistry.
ProteasePull-downwithProteaseInhibitor-coupledSepharose4B—Theproteaseinhibitorsaprotinin(5mg/mlSepharose),leupeptin(5mg/mlSepharose),andsoybeantrypsin
inhibitor(5mg/mlSepharose)wereimmobilizedtoCNBractivatedSepharose
TM 4B(GEHealthcare),asspecifiedbythe
manufacturer’sinstructions.Caco-2cellswerebiotin-labeled
andbiotinylatedproteinswerepulleddownwithmonomeric
avidinagarose.Thebiotinylatedproteinsweregentlyeluted
with4mMbiotinin50mMTris-HCl,pH8.5.Thebiotin-eluatewasseparatedfromtheavidin-agaroseandincubatedwith
60 lproteaseinhibitor-coupledSepharosein50mMTris-
HCl,pH8.75at37°Cfor30min.TheinhibitorSepharosewas
washed3with50mMTris-HCl,pH6.5.Proteaseswere
elutedfromtheinhibitor-coupledSepharoseusing0.1 Mglycine,pH2.4.Sampleswereneutralizedwith1MTrisimmediatelyafterelution.TheeluateswereaddedtoSDSsample
bufferandanalyzedbyWesternblotting.
GelatinZymography—Biotinylatedmonomericavidinagarose-purifiedproteinswereseparatedona7%SDS-polyacrylamidegelcontaining0.1%gelatin.Theproteinswerere-naturedbywashingthegelatingel230minin2.5%Triton
X-100inH 2 O.TowashoutexcessTritonX-100,thegelwas
washed210mininH 2 Oandthereafterincubatedin50mM
Tris-HCl,pH8.75overnightat37°C.Gelatinolyticbandson
thegelwerevisualizedbyCoomassieBrilliantBluestaining.
Immunofluorescence—Caco-2cellsgrownonfilterswere
fixedfor20minin4%paraformaldehydeinPBS(Bie&Berntsen)atroomtemperature.Thefollowingwasperformedat
4°C.Cellswerepermeabilizedwith0.05%TritonX-100in
PBSfor20min.UnspecificstainingwasblockedwithPBS
containing3%BSA(PBS/BSA)for30min.CellswereincubatedwithprimaryantibodydilutedinPBS/BSAfor1.5h,
washed3inPBSfollowedbyincubationwithrelevantAlexa
Fluor-conjugatedsecondaryantibodies(Invitrogen)for1h.
Whereindicatedthenucleiofthecellswerevisualizedby
4,6-diamidino-2-phenylindole(DAPI)staining.Thecells
werefinallymountedwithProlongGoldmountingmedium
(Invitrogen)andsubjectedtolaserscanningconfocalmicroscopyusingtheLeicaTCSSP2systemandZeissLS700system.
RESULTS
Steady-stateDistributionofMatriptase,HAI-1,and
Prostasin—Whereinthecellmatriptasecleavesitssubstrates
isnotwellunderstood,howeveritisfundamentaltounderstandinghowmatriptasemaintainsepithelialintegrity.Traffickingandactivationofendogenousmatriptase,prostasin,
andHAI-1wasthereforestudiedinCaco-2cells-ahuman
colonepithelialcellline.Uponreachingconfluence,Caco-2
cellsspontaneouslydifferentiateintoatightmonolayerof
polarizedcellswithanapicalandabasolateralplasmamembrane,separatedbytightjunctions.Matriptaseexpression
increasesduringCaco-2differentiation(2)consistentwiththe
higherlevelsofmatriptaseattheintestinalvilloustip(20).
Theamountofmatriptase-HAI-1complexalsoincreasesduringdifferentiationandreachesaplateauaroundday7–10in
differentiatingCaco-2cells(datanotshown).Forthatreason,
Caco-2cellsatday11post-confluencewereusedforthefollowingexperiments.
Wefirstinvestigatedthesteadystatedistributionof
matriptase,HAI-1,andprostasinbetweentheapicalandthe
basolateralplasmamembranedomains.Surfacebiotinylation
experimentsshowedthatthemajorityofmatriptasewaspresentamongbasolateralmembraneproteinsandmostprevalent
ina70kDaformcorrespondingtotheextracellulardomainof
matriptasecleavedatGly-149(Fig.1,lanes1and2),representingeitherzymogenmatriptaseorenzymaticallyactive
matriptase.Asmallfractionofthebasolateralmatriptasewas
detectedina120kDaform(Fig.1,lane2).The120kDaform
wasidentifiedasmatriptase-HAI-1complexbydetectionwith
boththeM69andanti-HAI-1antibodies(Fig.1,lanes4and
6).Twomatriptaseformsbelowthe120kDacomplexwere
alsodetectedonthebasolateralplasmamembrane(Fig.1,
lanes2and4).Thiscouldpossiblybedegradationproductsof
thematriptase-HAI-1complexoractivatedmatriptasein
complexwithotherinhibitors(21).
Full-lengthHAI-1hasanestimatedmolecularweightof55
kDaandwasdetectedonboththeapicalandthebasolateral
plasmamembrane(Fig.1,lanes5and6).Apicallylocated
HAI-1displayedahighermobilitythanbasolaterallylocated
HAI-1inthepresenceofSDS(Fig.1,lanes5and6)butthe
samemobilitywhenthesampleswereboiled(Fig.1,lanes7
and8).ThiscouldindicateaconformationaldifferencemakingapicalHAI-1moreresistanttodenaturationwithSDS.
Furtherstudiesareneededtoelucidatethenatureofthetwo
forms.AHAI-1complexaround85kDawasalsodetectedon
thebasolateralplasmamembrane(Fig.1,lane6).An85kDa
HAI-1complexwithmatriptaseaswellasprostasinhaspreviouslybeenreported(22,23).
Prostasinwaslocatedmainlyontheapicalplasmamembranealthoughminoramountscouldbedetectedonthebasolateralplasmamembrane(Fig.1,lanes9and10).These
dataareconsistentwiththeexistingliteratureshowingthe
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Matriptase Activates Prostasin on the Basolateral Plasma Membrane
FIGURE 1. Matriptase is located on the basolateral membrane whereas
prostasin is apically located. Caco-2 cells grown on Transwell filters were
biotin-labeled from either the apical (A) (lanes 1, 3, 5, 7, and 9) or basolateral
(B) side (lanes 2, 4, 6, 8, and 10). Biotinylated proteins were precipitated with
monomeric avidin, separated by SDS-PAGE and analyzed by Western blot
with antibodies against total matriptase (M32) (lanes 1 and 2), matriptase-
HAI-1 complex (M69) (lanes 3 and 4), the inhibitor HAI-1 (lanes 5– 8) and
prostasin (lanes 9 and 10) / boiling of samples. Samples analyzed with
the anti-prostasin antibody were boiled and reduced with DTT. The positions
of molecular weight markers are indicated on the left. Protein and
complex are marked with arrows and size on the right. The majority of the
plasma membrane-bound matriptase was located on the basolateral membrane
in a 70 kDa form, as detected with the antibody M32 (lane 2). A small
fraction of the basolateral plasma membrane-bound matriptase was found
in a 120 kDa form (lane 2). This form was also detected by the M69 (lane 4)
and anti-HAI-1 (lane 6) antibodies. HAI-1 was found both on apical and basolateral
plasma membrane (lanes 5– 8). Apical HAI-1 (lane 5) migrated
faster on the SDS-PAGE than the basolateral HAI-1 (lane 6) when samples
were not boiled but displayed the same size after boiling of the samples
(lanes 7 and 8). Plasma membrane-bound prostasin was located mainly on
the apical side (lane 9), although minor amounts were detected on the basolateral
membrane (lane 10). Results shown are representative of five independent
experiments.
basolaterallocalizationofmatriptaseandapicallocalizationof
prostasininapolarizedcell(15,18).
Next,weinvestigatedthesubcellularsteadystatedistributionofmatriptaseandHAI-1inCaco-2cellsbyimmunocytochemistry(Fig.2).Caco-2cellsweregrownonTranswellfiltersbeforefixation,permeabilization,andimmunolabeling.
MatriptaseandHAI-1werebothdetectedonthebasolateral
plasmamembrane(Fig.2,A–F).Interestingly,matriptaseand
HAI-1werealsodetectedinstructuresneartheapicalplasma
membrane(Fig.2F).Onlyweakdetectionofmatriptase-
HAI-1complexwasobservedonthebasolateralplasmamem-
brane(Fig.2D).Surprisinglythemajorityofthematriptase-
HAI-1complexwasdetectedinstructuresneartheapical
plasmamembrane(Fig.2,D–F).Tofurtherinvestigatethe
locationofthematriptase-HAI-1complexintheapicalregion
ofthecells,Caco-2cellswereimmunolabeledwithandwithoutpermeabilizationoftheplasmamembrane(Fig.2,Gand
H).Withoutpermeabilization,almostnomatriptase-HAI-1
complexcouldbedetected(Fig.2H),however,withpermeabilizationadistinctlabelingofthevesicularstructureswas
clearlyvisiblebelowtheapicalplasmamembrane(Fig.2G),
confirminganintracellularlocalizationofthestructurescontainingthematriptase-HAI-1complex.Thesedataareconsistentwithourbiotinylationexperiments,aswewereonly
abletolabelmatriptaseonthebasolateralplasmamembrane
(Fig.1,lanes1and3).
TheMatriptase-HAI-1ComplexIsGeneratedduring
Endocytosis—Oursteadystateexperimentsshowthatalarge
portionofmatriptaseispresentina70kDaformonthebasolateralplasmamembrane.Furthermore,theexperiments
showthatlargeamountsofmatriptase-HAI-1complexare
detectedinintracellularstructures.Thissuggeststhat
matriptaseisendocytosedfromtheplasmamembraneand
inhibitedbyHAI-1duringthisprocess.
WeexaminedtheendocytosisofbasolateralplasmamembraneboundmatriptaseandHAI-1usingbiotinylationand
internalizationtechniques(see“ExperimentalProcedures”).
Theendocytosisexperimentsshowedthatmatriptaseisvery
efficiently,andalmostcompletely,endocytosedwithin60min
fromthebasolateralmembrane(Fig.3A).Furthermore,a
3-foldincreaseinthesignalofmatriptase-HAI-1complexes
wasdetectedwithinthefirst90minofincubation,showing
thatthe70kDamatriptaseformsacomplexwithHAI-1duringtheendocytosis(Fig.3B).Severalmatriptase-HAI-1complexesbetween95and120kDaweregeneratedfromthe70
kDaformduringtheendocytosis,asdetectedbyboththe
matriptaseandHAI-1antibodies(Fig.3,A–C).After90min,
allofthesurface-labeledmatriptasewasfoundincomplex
withHAI-1andwaslocatedexclusivelyintracellularly
(Fig.3B).
The55kDaHAI-1hadanendocytosispatterndifferent
fromHAI-1incomplexwithmatriptase.The55kDaHAI-1
wasonlypartiallyendocytosed,withthelargestintracellular
poolseenafter15min.ThissuggeststhatHAI-1,whennotin
complexwithmatriptase,isrecyclingtotheplasmamembranefromearlyendosomes(Fig.3C).Thistypeofrecycling
ofHAI-1hasbeenshownpreviouslyinMDCKcells(24).
AHAI-1complexaround85kDawasobservedintheTotal
panelfromtimepoint0to15min,whichwasnotdetectedby
thematriptaseantibodies(Fig.3,compareCtoAandB,respectively).ThiscouldpossiblybeaHAI-1-prostasincomplex.Asimilarcomplexhaspreviouslybeenreportedinkeratinocytes(22).Togetherthesedatashowthatmatriptaseis
efficientlyendocytosedfromthebasolateralsideandformsa
complexwithHAI-1duringthisprocess.
MatriptaseZymogenIsCleavedtotheActiveProteaseonthe
SurfaceoftheBasolateralPlasmaMembrane—Wehaveupto
nowshowedthatmatriptaseispresentpredominantlyina70
kDaformonthebasolateralplasmamembraneandisefficientlyendocytosedformingacomplexwithHAI-1thataccumulatesinintracellularstructures,makingitaprerequisite
thatmatriptaseactivationoccurspriortoendocytosis.
Whethermatriptaseactivationoccursbeforearrivaloratthe
plasmamembraneisnotwelldefined.
Weexaminedtheactivationcleavageofbasolateralplasma
membraneboundmatriptaseusingsurfacebiotinylationand
anincubationassay.Underreducingconditions,matriptase
zymogenisa70kDaproteinwhileenzymatically-active
matriptaseseparatesintotwofragments;thestemdomain
andthe30kDaproteasedomain.TheantibodyIM1014reacts
withtheserineproteasedomainofbothzymogenandactivatedmatriptaseunderreducingconditions.Thisexperiment
showedthatminoramountsofthe70kDamatriptasezymogenwerepresentonthebasolateralplasmamembranetogetherwith30kDacleavedmatriptaseserineproteasedomain
(Fig.4,lane1).Thebasolateralmatriptasezymogenwasrapidlycleavedasonlythe30kDabandcouldbedetectedafter
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Matriptase Activates Prostasin on the Basolateral Plasma Membrane
FIGURE 2. Matriptase-HAI-1 complex accumulates in intracellular structures. Caco-2 cells grown on Transwell filters, were fixed, permeabilized, and immunolabeled
with antibodies against total matriptase (M32) and HAI-1 (A–C) or matriptase-HAI-1 complex (M69) and HAI-1 (D–F). Using a confocal scanning
microscope, images were taken in the XY plane showing a single section through the monolayer and the XZ plane showing a cross section of the monolayer.
The position of the plane of the XY section is indicated on the XZ plane (black arrows) on the right. The scale bar represents 10 m. A–C, Matriptase
was detected on the basolateral plasma membrane of the Caco-2 cells co-localizing with HAI-1. Matriptase and HAI-1 was also co-localizing in structures
near the apical membrane (white arrowheads). D–F, matriptase-HAI-1 complex was observed in structures near the apical plasma membrane (white arrowheads).
Low amounts of matriptase-HAI-1 complex were detected on the basolateral plasma membrane. HAI-1 was detected both on apical and basolateral
plasma membranes as well as in the apical structures co-localizing with matriptase. G and H, Caco-2 cells were treated with or without Triton X-100 prior to
immunolabeling with the antibody M69. G, in the cells permeabilized with Triton X-100, a distinct detection of matriptase-HAI-1 complex was observed in
apical vesicular structures. H, in cells without permeabilization, the vesicular structures with matriptase-HAI-1 complex were not detected. Only a
weak basolateral signal was observed. The nuclei were visualized by DAPI staining shown in blue. Results shown are representative of three independent
experiments.
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just5minofincubation(Fig.4,lane2).Aslightincreasein
the30kDabandintensitywasobservedupto15minafter
labeling,mostlikelycausedbyahigheraffinityofIM1014
antibodyforthecleaved30kDaproteasedomainthanthe70
kDaform.Thesedatasuggestthatmatriptaseisfastandefficientlycleavedtoitsactiveformonthebasolateralcell
surface.
ActiveMatriptaseIsPresentontheBasolateralPlasma
Membrane—Ithaspreviouslybeenreportedthattheperiodof
timeforactivematriptasetoactonitssubstratesisverylimitedasmatriptaseactivationistightlycoupledtoinhibitionby
HAI-1.Thiswindowofactionisassumedtobeinbetweenthe
proteolyticcleavagecreatingthefullyactiveproteaseandthe
rapidcomplexformationwithitsinhibitorHAI-1.Ourpreviousexperimentssuggestthatbothmatriptaseactivationand
inhibitiontakesplaceonthebasolateralmembraneandwe
wouldthereforeexpecttofindfreeactivematriptaseonthe
basolateralplasmamembrane.Toaddressthis,weinvestigatedwhethermatriptaseatthebasolateralplasmamembrane
wasabletobindtothegeneralserineproteaseinhibitors,
aprotininandleupeptin,towhichweexpectonlytheactive
proteasetobind.Wewereabletopurifymatriptasefromthe
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Matriptase Activates Prostasin on the Basolateral Plasma Membrane
FIGURE 3. Matriptase in complex with HAI-1 is generated during the efficient endocytosis of matriptase from the basolateral plasma membrane.
Caco-2 cells were grown on Transwell filters, and proteins on the basolateral plasma membrane were biotinylated at 4 °C using a cleavable biotinylation
reagent, s-NHS-SS-biotin. The labeled cells were incubated at 37 °C for the time indicated (0 –120 min). After incubation, the proteins remaining on the
plasma membrane were biotin-stripped, using the membrane non-permeant reducing agent glutathione, leaving only endocytosed proteins biotinylated
(endocytosis panels). Biotin reduction was omitted in a parallel set of samples to monitor the degradation of the biotinylated proteins over time (total panels).
Biotinylated proteins were precipitated with monomeric avidin agarose, and the avidin pull-downs were analyzed with SDS-PAGE and Western blotting
using the antibodies (A) M32 against total matriptase, (B) M69 against matriptase-HAI-1 complex, and (C) M19 against HAI-1. The positions of molecular
markers (kDa) are indicated on the left. Proteins and complexes are indicated with arrows and size. Quantification of bands from Western blots was done
using the software ImageJ. A graphic presentation of three independent endocytosis experiments is shown on the right hand side. The dotted line equals
total protein and solid line equals endocytosed protein. The standard deviation is shown with error bars. A, matriptase was endocytosed from the basolateral
plasma membrane within 60 min. Approximately 80% of matriptase was endocytosed at 60 min as indicated on the graph. B, during the 120-min incubation
there was a 3-fold increase in the matriptase-HAI-1 complexs. C, free 55 kDa HAI-1 was partially endocytosed within 15 min. (C, endocytosis panel).
Results shown are representative of three independent experiments.
basolateralplasmamembrane,withbothaprotinin-andleupeptin-coupledSepharose(Fig.5A,lanes3and5).No
matriptasewaspurifiedwiththenegativecontrolsoybean
trypsininhibitor-SepharoseoruncoupledSepharose(datanot
shown).Theinhibitor-boundfractionofmatriptasewasnot
detectedbytheantibodydetectingmatriptase-HAI-1complexes(Fig.5B,lanes3and5).
Wewantedtoverifythatthepurifiedmatriptasedetected
usingtheinhibitor-coupledSepharoseindeedwasfreeactive
matriptaseandnotactivatedmatriptasedissociatedfromthe
matriptase-HAI-1complexes.Totestthis,asamplecontainingmostlymatriptase-HAI-1complexwasexposedtothe
inhibitor-coupledSepharose.Nodetectablematriptasefrom
thematriptase-HAI-1complexwaspurifiedwiththeinhibitor
Sepharose(Fig.5,lanes4and6).Thisverifiesthatitisfree
activematriptasethatisbeingpulleddownandthatthe
matriptase-HAI-1complexisnotdissociatedduringourcell
extractionprocedure.
Finally,totesttheproteolyticactivityofmatriptase,thetwo
biotinylatedfractions,containing70kDamatriptaseand
matriptase-HAI-1complexes,respectively,wereanalyzedfor
theirabilitytodisplaygelatinolyticactivityatpH8.75,where
matriptasehasoptimalenzymaticactivity(25)(Fig.5C).The
fractionwiththe70kDamatriptasedisplayedonegelatinolyticbandaround70kDaatpH8.75,correspondingtothe
sizeoffreeactivematriptase(Fig.5C,lane1).Thefraction
containingmostlymatriptase-HAI-1complex,showedonly
weakgelatinolyticactivityaround70kDa(Fig.5C,lane2).
Thus,thegelatinolyticactivitypatternmatchestheoneobservedformatriptaseimmunoreactivitywiththeantibody
M32(Fig.5A)implyingthatthegelatinolyticactivityobserved
iscausedbymatriptase.
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5798 JOURNAL OF BIOLOGICAL CHEMISTRY VOLUME 286 • NUMBER 7 • FEBRUARY 18, 2011

Matriptase Activates Prostasin on the Basolateral Plasma Membrane
FIGURE 4. Matriptase is cleaved to the two chain form on the basolateral
cell surface. Caco-2 cells were grown on Transwell filters, biotinylated
from the basolateral side at 4 °C and incubated up to 60 min at 37 °C after
labeling. Biotinylated proteins were pulled down with streptavidin-agarose,
boiled, reduced and analyzed by SDS-PAGE and Western blotting using the
antibody IM1014. The positions of molecular markers (kDa) are indicated on
the left. Bands are indicated with arrows and size. At time 0 the matriptase is
detected as a 70 kDa band, representing the non-cleaved zymogen and a
30 kDa band representing the cleaved protease domain (lane 1). After 5 min
of incubation the zymogen could no longer be detected and an increase in
the 30 kDa serine protease domain was detected (lanes 2– 6). Results shown
are representative of two independent experiments.
FIGURE 5. Matriptase is active on the basolateral plasma membrane
but not intracellularly. Caco-2 cells on Transwell filters were surfacebiotinylated
from the basolateral side at 4 °C. Some were lysed immediately
after biotinylation (0) and some were incubated for 2 h at 37 °C to
transform all biotin-labeled matriptase into activated matriptase in complex
with HAI-1 (2). Biotinylated proteins were precipitated with monomeric
avidin and gently eluted with biotin. The avidin pull-downs were
divided into three groups: No further treatment (lanes 1 and 2), pulldown
with aprotinin-Sepharose (lanes 3 and 4) and pull-down with leupeptin-Sepharose
(lanes 5 and 6). The samples were analyzed by SDS-
PAGE and Western blot with antibodies against total matriptase (M32)
and matriptase-HAI-1 complex (M69). A, only at time 0 was it possible to
pull-down M32-detectable matriptase with both aprotinin and leupeptin.
B, M69-detectable matriptase was pulled down with the monomeric
avidin (B, lanes 1 and 2), but was lost with additional inhibitor pull-down
(B, lanes 3– 6). C, two avidin-purified fractions showed gelatinolytic properties
with a band around 70 kDa (C, lanes 1 and 2), matching the size
and pattern of A, lanes 1 and 2. Results shown are representative of two
independent experiments.
FIGURE 6. Active prostasin is present on the basolateral as well as on
the apical plasma membrane. Caco-2 cells on Transwell filters were surface
biotinylated from either the apical or basolateral side at 4 °C. Biotinylated
proteins were precipitated with monomeric avidin and gently eluted
with biotin. The avidin pull-downs were divided into three: No further treatment
(lanes 1 and 2), pull-down with aprotinin-Sepharose (lanes 3 and 4)
and pull-down with trypsin inhibitor as negative control (lanes 5 and 6). The
pull-down fractions were analyzed by SDS-PAGE and Western blotting.
Prostasin was purified from both apical plasma membrane (lane 1) and basolateral
plasma membrane (lane 2). The majority of the apical prostasin
could be pulled down with the serine protease inhibitor aprotinin (lane 3). A
small but significant fraction of basolateral prostasin was also able to bind
aprotinin and hence was in its active form (lane 4). No prostasin binding was
observed when using the negative control trypsin-coupled Sepharose
(lanes 5 and 6). Results shown are representative of three independent
experiments.
ActiveProstasinIsPresentonBoththeApicalandtheBasolateralPlasmaMembrane—Allofourdatasuggestthat
matriptaseisactiveandabletocleaverelevantsubstratesina
shortperiodoftimeonthebasolateralplasmamembranebeforeitformsacomplexwithHAI-1andisendocytosed.If
prostasinisindeedasubstrateformatriptaseinpolarizedepithelialcells,wewouldexpecttofindthecleavedformofprostasinonthebasolateralmembrane.Totestthis,weutilized
thefactthatprostasinisaserineproteaseandonlythe
cleavedformisproteolyticallyactiveandtherebyabletobind
serineproteaseinhibitorssuchasaprotinin.Ourexperiment
showedthatprostasinonboththeapicalandthebasolateral
plasmamembranewasabletobindtoaprotinin(Fig.6,lanes
3and4).Thissuggeststhatprostasinisactiveonboththe
apicalandthebasolateralsideofthecell.
MatriptaseCo-localizeswithProstasinontheBasolateral
PlasmaMembrane,SubsequentlyProstasinIsTranscytosedto
theApicalPlasmaMembrane—Weknowfromthesurface
labelingexperimentthatthemajorityofmembrane-bound
prostasinatsteadystateislocatedontheapicalside(Fig.1,
lane9)whereasonlyaminorfractionislocatedonthebasolateralmembraneinCaco-2cells(Fig.1,lane10).Fromour
experiments,wealsoknowthatprostasinisnotonlyfoundin
itsactiveformontheapicalplasmamembranebutalsoonthe
basolateralplasmamembrane.Thiscouldindicatethatprostasinistransportedtothebasolateralplasmamembraneto
getactivatedbymatriptase.
Wewantedtoinvestigatetheendocytictransportofthe
apicalandbasolateralprostasin.Initially,wetestedifthe
twomembranefractionswereendocytosedusingthesame
experimentalsetupasformatriptaseandHAI-1endocytosis.
Wefoundthatprostasinwasnotendocytosedfromtheapical
plasmamembranebutwascompletelyendocytosedfromthe
basolateralplasmamembranewithin60min(datanot
shown).Thismadeusquestionifprostasinisinitiallytransportedtothebasolateralmembraneandactivatedby
matriptasebeforeitisre-routedbytranscytosistotheapical
membrane.Alongresidencetimeattheapicalplasmamem-
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FEBRUARY 18, 2011 • VOLUME 286 • NUMBER 7 JOURNAL OF BIOLOGICAL CHEMISTRY 5799

Matriptase Activates Prostasin on the Basolateral Plasma Membrane
FIGURE 7. Prostasin and HAI-1 are transcytosed from the basolateral to the apical plasma membrane. Caco-2 cells were surface biotinylated from the
basolateral side at 4 °C. The labeled cells were incubated at 37 °C for the time indicated in the internalization panel (4, 12, or 18 h as indicated). After incubation
the proteins remaining on the plasma membrane were biotin-stripped, using the membrane non-permeant reducing agent glutathione, from either
apical (A red.), basolateral (B red.), or both sides (A B red.). Biotin reduction was omitted in a set of samples to monitor the total amount of biotinylated
protein remaining after the incubation (Total). The experiment was performed in duplicates. Biotinylated proteins were precipitated with monomeric avidin
agarose, separated with SDS-PAGE and analyzed by Western blotting. A, prostasin could be biotin-stripped from the apical side (A red.) after 18 h of internalization,
as a decrease in signal was observed compared with the total samples. This shows that prostasin has moved from the basolateral to the apical side.
When surface proteins were biotin-stripped from the basolateral side no decrease in signal was observed (B red.), once again showing that no biotinylated
prostasin was left on the basolateral side after 18 h. To clarify if some of the prostasin was located intracellularly, the cells were biotin stripped from both
apical and basolateral side (A B red.). Biotin-stripping from both apical and basolateral side removed the majority of the biotinylated prostasin suggesting
that all of prostasin had within the 18 h transferred from the basolateral to the apical plasma membrane. B, HAI-1 was transcytosed from the basolateral to
the apical plasma membrane within 12 h. Most of HAI-1 could be biotin stripped from the apical side after incubation (A red.) where a major fraction was left
after basolateral reduction (B. red) showing that a large fraction of biotinylated HAI-1 had moved from the basolateral to the apical plasma membrane. It
was demonstrated that the faster migrating form of HAI-1 resides at the apical plasma membrane 12 h after biotinylation and the slower migrating form
remained at the basolateral plasma membrane, as these could be biotin-stripped from their respective sites (A red. compared with B red., respectively).
C, after just 4 h of incubation most biotinylated matriptase was no longer detectable in the cell. The remaining matriptase was mainly found in the 120 kDa
complex with HAI-1. A fraction of the matriptase-HAI-1 complex could be biotin stripped from the basolateral side (B red.) but not the apical side (A red.).
D, M69 detectable matriptase showed the same amount of matriptase-HAI-1 complex at time 0 as for 4 h of incubation. A fraction of the complex was reducible
from the basolateral side after 4 h of incubation (B red.). Results shown are representative of three independent experiments.
branecombinedwithashortresidencetimeatthebasolateral
plasmamembranewouldgiveasteadystateaccumulationat
theapicalplasmamembrane.We,therefore,testedifthebasolaterallyendocytosedfractionwastranscytosedtotheapical
membrane.
Transcytosisforseveralincubationtimeswasinvestigated
(1,2,4,8,12,and18h)ThetimepointsmostclearlydemonstratingtranscytosisareshowninFig.7.Transcytosisofprostasinwasdetectableafter8h(datanotshown)andmostof
prostasinwastranscytosedfromthebasolateraltotheapical
plasmamembraneafter18h(Fig.7A).HAI-1wasalsotranscytosed,however,lessefficientlythanprostasin(Fig.7B).
Ourresultsstronglysuggestthattheslowmigratingformof
HAI-1atthebasolateralplasmamembraneistheprecursor
forthefastermigratingHAI-1presentontheapicalplasma
membrane(Fig.7B).
Matriptase-HAI-1 complex has been purified from milk,
suggesting that matriptase is secreted from the apical
plasma membrane (16). However, we were unable to detect
transcytosis of matriptase from the basolateral to the apical
plasma membrane, as matriptase has a shorter half-life in
the cell than HAI-1 and prostasin (Fig. 7, C and D). The
remaining biotin-labeled matriptase was found in the
matriptase-HAI-1 complexes between 95 and 120 kDa after
4 h incubation (Fig. 7, C and D). These complexes were
also detected with the anti-HAI-1 antibody after 4 h incubation
(data not shown). Instead, basolaterally biotin-labeled
matriptase could be detected in the basolateral media
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5800 JOURNAL OF BIOLOGICAL CHEMISTRY VOLUME 286 • NUMBER 7 • FEBRUARY 18, 2011

Matriptase Activates Prostasin on the Basolateral Plasma Membrane
A
B
C
D
Total matriptase
(M32)
Matriptase-HAI-1
complex (M69)
HAI-1
Prostasin
5m 15m 30m 1h 2h 4h 8h 18h
- 120 kD
Apical media
- 70 kD
- 120 kD
- 70 kD
- 120 kD
- 50 kD
- 40 kD
FIGURE 8. Matriptase is shed into the basolateral media while basolateral
prostasin is transcytosed and shed into the apical media. Caco-2
cells were surface-biotinylated from the basolateral side at 4 °C. Cells were
incubated at 37 °C after labeling and apical and basolateral media was collected
at 5, 15, 30 min, 1, 2, 4, 8, and 18 h to reveal any shedding of the biotin-labeled
matriptase, HAI-1 and prostasin. Biotinylated proteins from the
media were pulled down with monomeric avidin agarose, separated with
SDS-PAGE and analyzed by Western blotting. A, matriptase was released to
the basolateral media within 1 h as three forms: a 70 kDa form and two proteolytically
shed complexes at 85 and 110 kDa. B, two complexes of size 85
and 110 kDa could be detected with the M69 antibody within 1–2 h, suggesting
these to be matriptase-HAI-1 complexes. C, no free 55 kDa HAI-1
was found in the media, only HAI-1 in the two complexes at 85 and 110 kDa
was detected in the basolateral media after 1 h and accumulated up to 18 h.
D, basolateral prostasin was detected in the apical media after 18 h incubation,
confirming transcytosis before secretion into the apical media. Results
shown are representative of two independent experiments.
after only 1 h and matriptase accumulated in the media
over the 18 h (Fig. 8, A and B). Both the 70 kDa form as
well as two forms of the matriptase-HAI-1 complex around
110 and 85 kDa were detected in the media. Only HAI-1 in
complex with matriptase was detected in the basolateral
media (Fig. 8C), whereas no free HAI-1 could be detected.
None of the biotin-labeled matriptase or HAI-1 could be
detected in the apical media (data not shown). Basolateral
prostasin could not be detected in the basolateral media
(data not shown) but after 18 h of incubation after the biotinylation,
prostasin appeared in the apical media (Fig. 8D).
This is consistent with the finding that prostasin is transcytosed
from the basolateral to the apical plasma membrane
within this time frame. Thus, we were able to show that
prostasin on the basolateral plasma membrane is transcytosed
to the apical plasma membrane from where it is shed
to the media. This suggests that prostasin is routed via the
basolateral plasma membrane where it is activated before it
is transcytosed to the apical membrane, thereby providing
a mechanism for activation of the prostasin zymogen in
polarized epithelial cells by matriptase or other basolaterally
located serine proteases, despite the separate steadystate
localization of the proteases.
DISCUSSION
Ithasbeenshownthatmatriptaseandprostasinareconstitutivelyexpressedandco-localizeinmostepitheliaincluding
thetissuesaffectedbymatriptaseablation,suggestingapossibleglobalroleforamatriptase-prostasincascadeinepithelial
homeostasis(26).Inapolarizedcell,matriptaseisatsteadystateconcentratedandlocalizedmainlyatthebasolateral
plasmamembranetogetherwithHAI-1(15,24)whileprostasinismainlyconcentratedandlocalizedtotheapicalplasma
membrane(17,18).Ourdataprovideapossiblesolutionto
howmatriptasecanactivateprostasindespitetheirtwodifferentsubcellularlocalizations,byshowingthatthetwomoleculesmeetenroute.
Matriptaseismainlylocatedonthebasolateralplasma
membranewhereitisactivated,inhibited,endocytosedin
complexwithitsinhibitorHAI-1andisaccumulatedinintracellularstructures.ObservationofthetransmembraneNterminalfragmentofmatriptaseinintracellularcompartmentshaspreviouslybeendescribedinratenterocytes(14).
Weshowherethatprostasinismainlylocatedontheapical
plasmamembraneinitsactiveform.Interestingly,wefinda
smallfractionofprostasinonthebasolateralplasmamembraneco-localizingwithmatriptase,makingitpossiblefor
proteaseandsubstratetointeractandactivationtooccur.In
agreementwiththis,weshowthatpartofthebasolateral
prostasinisactivebyitsabilitytobindtogeneralserineproteaseinhibitors.Thebasolateralprostasinisendocytosedand
transcytosedtotheapicalplasmamembrane,whereit
accumulates.
Bothmatriptaseandprostasinhasbeenshowntobeableto
activatetheepithelialsodiumchannel(ENaC)inXenopus
oocytes(5,27,28).ENacisanepithelialmembrane-bound
sodiumchannellocatedintheapicalmembraneofpolarized
cellsandisrequiredfornormalepidermaldifferentiation(29,
30).Becausematriptaseistargetedtothebasolateralmembranewhereitisactivatedandrapidlyinhibiteditismore
likelythatprostasinwhichco-localizeswithENaContheapicalmembraneisacandidateactivatorinpolarizedcells.Both
matriptaseandprostasinhavebeencoupledtomaintenance
offunctionaltightjunctionsinavarietyofepithelialcelltypes.
InaCaco-2model,lossofmatriptasewasassociatedwithenhancedexpressionandincorporationofthepore-forming
proteinclaudin-2attightjunctions(2).Prostasinhasbeen
reportedtoregulatetightjunctions,paracellularpermeability
andTEERbyaproteaseactivity-dependentmechanismin
renalcells(4,5).Thismeansthatthetwoproteasesareboth
involvedincascadesimportantfortightjunctionformationin
polarizedepithelia.
Increasingevidencesuggeststhatserineproteasescontributeinacomplexwaytotheregulationofintestinalintegrity
andbarrierfunction.Proteaseinhibitorshavebeenshownto
suppresstheformationoftightjunctionsingastrointestinal
celllinessuggestingthatproteolyticactivityisnecessaryfora
functionalepithelialbarrier(31).Theepithelialbarrieriscrucialinthegastrointestinaltractandisoftencompromisedin
inflammatoryboweldiseaseslikeCrohndiseaseandulcerativecolitis.ThesodiumchannelENaCisdown-regulatedin
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FEBRUARY 18, 2011 • VOLUME 286 • NUMBER 7 JOURNAL OF BIOLOGICAL CHEMISTRY 5801

Matriptase Activates Prostasin on the Basolateral Plasma Membrane
inflammatoryboweldiseasesandhasbeenproposedasatherapeutictarget(32–34).Itwillbeimportantforfuturestudies
toidentifytheroleofthematriptase-prostasincascadeand
thesubstratesinvolvedinthispathwaytoinvestigatetherole
ofmatriptaseininflammatorydiseasesofthegastrointestinal
tract.
Together,ourdatashowthatactivematriptaseanditssubstrateprostasinco-localizeatthebasolateralplasmamembraneandherebyprovideavenueforamatriptase-prostasin
zymogencascadetobeinitiatedinpolarizedepithelialcells.
Furthermore,weshowhowprostasinistransportedafteractivationtotheapicalmembranetoco-localizewithitsprimary
substrateENaC.
Acknowledgment—WethankDr.MaryJoDantonforcritically
readingthemanuscript.
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5802 JOURNAL OF BIOLOGICAL CHEMISTRY VOLUME 286 • NUMBER 7 • FEBRUARY 18, 2011

Paper II
Reduced prostasin (CAP1/PRSS8) activity eliminates HAI-1 and HAI-2 deficiency-associated
developmental defects by preventing matriptase activation
Roman Szabo 1 , Katiuchia Uzzun Sales 1 , Peter Kosa 1 , Natalia A. Shylo 1 , Sine Godiksen 1,2,3 ,
Karina K. Hansen 1 , Stine Friis 1 , J. Silvio Gutkind 1 , Lotte K. Vogel 2 , Edith Hummler 4 , Eric
Camerer 5,6 , and Thomas H. Bugge 1
1 Oral and Pharyngeal Cancer Branch, National Institute of Dental and Craniofacial Research,
National Institutes of Health, Bethesda, MD, USA.
2 Department of Cellular and Molecular Medicine, University of Copenhagen, Copenhagen,
Denmark.
3 Department of Biology, University of Copenhagen, Copenhagen, Denmark.
4 Pharmacology and Toxicology Department, University de Lausanne, Lausanne, Switzerland.
5 INSERM U970, Paris Cardiovascular Research Centre, Paris, F-75015, France. 6Université Paris-
Descartes, Paris, F-75006, France.
Published in PLoS Genetic, August 2012
47

Reduced Prostasin (CAP1/PRSS8) Activity Eliminates HAI-
1 and HAI-2 Deficiency–Associated Developmental
Defects by Preventing Matriptase Activation
Roman Szabo 1 , Katiuchia Uzzun Sales 1 , Peter Kosa 1 , Natalia A. Shylo 1 , Sine Godiksen 1,2,3 ,
Karina K. Hansen 1 , Stine Friis 1 , J. Silvio Gutkind 1 , Lotte K. Vogel 2 , Edith Hummler 4 , Eric Camerer 5,6 ,
Thomas H. Bugge 1 *
1 Oral and Pharyngeal Cancer Branch, National Institute of Dental and Craniofacial Research, National Institutes of Health, Bethesda, Maryland, United States of America,
2 Department of Cellular and Molecular Medicine, Faculty of Health Science, University of Copenhagen, Copenhagen, Denmark, 3 Department of Biology, Faculty of
Science, University of Copenhagen, Copenhagen, Denmark, 4 Pharmacology and Toxicology Department, University de Lausanne, Lausanne, Switzerland, 5 INSERM U970,
Paris Cardiovascular Research Centre, Paris, France, 6 Université Paris-Descartes, Paris, France
Abstract
Loss of either hepatocyte growth factor activator inhibitor (HAI)-1 or -2 is associated with embryonic lethality in mice, which
can be rescued by the simultaneous inactivation of the membrane-anchored serine protease, matriptase, thereby
demonstrating that a matriptase-dependent proteolytic pathway is a critical developmental target for both protease
inhibitors. Here, we performed a genetic epistasis analysis to identify additional components of this pathway by generating
mice with combined deficiency in either HAI-1 or HAI-2, along with genes encoding developmentally co-expressed
candidate matriptase targets, and screening for the rescue of embryonic development. Hypomorphic mutations in Prss8,
encoding the GPI-anchored serine protease, prostasin (CAP1, PRSS8), restored placentation and normal development of
HAI-1–deficient embryos and prevented early embryonic lethality, mid-gestation lethality due to placental labyrinth failure,
and neural tube defects in HAI-2–deficient embryos. Inactivation of genes encoding c-Met, protease-activated receptor-2
(PAR-2), or the epithelial sodium channel (ENaC) alpha subunit all failed to rescue embryonic lethality, suggesting that
deregulated matriptase-prostasin activity causes developmental failure independent of aberrant c-Met and PAR-2 signaling
or impaired epithelial sodium transport. Furthermore, phenotypic analysis of PAR-1 and matriptase double-deficient
embryos suggests that the protease may not be critical for focal proteolytic activation of PAR-2 during neural tube closure.
Paradoxically, although matriptase auto-activates and is a well-established upstream epidermal activator of prostasin,
biochemical analysis of matriptase- and prostasin-deficient placental tissues revealed a requirement of prostasin for
conversion of the matriptase zymogen to active matriptase, whereas prostasin zymogen activation was matriptaseindependent.
Citation: Szabo R, Uzzun Sales K, Kosa P, Shylo NA, Godiksen S, et al. (2012) Reduced Prostasin (CAP1/PRSS8) Activity Eliminates HAI-1 and HAI-2 Deficiency–
Associated Developmental Defects by Preventing Matriptase Activation. PLoS Genet 8(8): e1002937. doi:10.1371/journal.pgen.1002937
Editor: Hamish S. Scott, SA Pathology, Australia
Received April 5, 2012; Accepted July 18, 2012; Published August 30, 2012
This is an open-access article, free of all copyright, and may be freely reproduced, distributed, transmitted, modified, built upon, or otherwise used by anyone for
any lawful purpose. The work is made available under the Creative Commons CC0 public domain dedication.
Funding: The study was supported by the NIDCR Intramural Research Program (THB), the Augustinus Foundation, Købmand Kristian Kjær og Hustrus
Foundation, the Kjær-Foundation, Dagmar Marshalls Foundation, Snedkermester Sophus Jacobsen og Hustru Astrid Jacobsens Foundation, Grosserer Valdemar
Foersom og Hustru Thyra Foersoms Foundation, Fabrikant Einar Willumsens Mindelegat, the Harboe Foundation (SG and LKV), the Lundbeck Foundation (KKH,
SG, and LKV), the Swiss National Science Foundation 31003A-127147/1 (EH), the INSERM Avenir, Marie Curie Actions, the French National Research Agency, and
the Ile-de-France Region (EC). The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.
Competing Interests: The authors have declared that no competing interests exist.
* E-mail: thomas.bugge@nih.gov
Introduction
Studies conducted within the past two decades have uncovered
a large family of membrane-anchored serine proteases that
regulates vertebrate development, tissue homeostasis, and tissue
repair by providing focal proteolysis essential for cytokine and
growth factor maturation, extracellular matrix remodeling,
signaling receptor activation, receptor shedding, regulation of
ion channel activity, and more (reviewed in [1,2,3]). Individual
members of this family regulate both vertebrate development and
postnatal tissue homeostasis, including auditory and vestibular
system development [4,5,6], differentiation of stratified epithelia
[7,8], loss of epithelial tight junction function [9,10], failure to
activate digestive enzymes [11], thyroid hormone availability [4],
sodium and water homeostasis [12,13,14], iron homeostasis
[15,16], and fertility [17,18]. Likewise, mounting evidence suggests
that excessive or spatially dysregulated membrane-anchored serine
protease activity contributes to several human disorders, including
congenital malformations [19], epithelial dysfunction [20,21,22],
and cancer [3].
Matriptase is a modular type II transmembrane serine protease,
encoded by the ST14 gene, that has pleiotropic functions in
epithelial development and postnatal homeostasis, at least in part
through its capacity to regulate epithelial tight junction formation
in simple and stratified epithelia [2,3]. In the human and mouse
epidermis, matriptase appears to function as part of a proteolytic
cascade in which it acts upstream of the GPI-anchored serine
protease prostasin (CAP1/PRSS8), most likely by directly activat-
PLOS Genetics | www.plosgenetics.org 1 August 2012 | Volume 8 | Issue 8 | e1002937

Embryonic Cell Surface Serine Protease Cascade
Author Summary
Vertebrate embryogenesis is dependent upon a series of
precisely coordinated cell proliferation, migration, and
differentiation events. Recently, the execution of these
events was shown to be guided in part by extracellular
cues provided by focal pericellular proteolysis by a newly
identified family of membrane-anchored serine proteases.
We now show that two of these membrane-anchored
serine proteases, prostasin and matriptase, constitute a
single proteolytic signaling cascade that is active at
multiple stages of development. Furthermore, we show
that failure to precisely regulate the enzymatic activity of
both prostasin and matriptase by two developmentally coexpressed
transmembrane serine protease inhibitors,
hepatocyte growth factor activator inhibitor-1 and -2,
causes an array of developmental defects, including clefting
of the embryonic ectoderm, lack of placental labyrinth
formation, and inability to close the neural tube. Our study
also provides evidence that the failure to regulate the
prostasin–matriptase cascade may derail morphogenesis
independent of the activation of known protease-regulated
developmental signaling pathways. Because hepatocyte
growth factor activator inhibitor–deficiency in humans is
known to cause an assortment of common and rare
developmental abnormalities, the aberrant activity of the
prostasin–matriptase cascade identified in our study may
contribute importantly to genetic as well as sporadic birth
defects in humans.
ing the prostasin zymogen [23,24,25,26]. Several additional
candidate proteolytic substrates have been identified for matriptase
in cell-based and biochemical assays, including growth factor
precursors [27,28,29,30], protease-activated signaling receptors
[31,32,33], ion channels [34,35], and other protease zymogens
besides pro-prostasin [29,36,37]. However, the extent to which
cleavage of these substrates is critical to matriptase-dependent
epithelial development and maintenance of epithelial homeostasis
needs to be established.
Although matriptase is not required for term development in
humans and most mouse strains ([24,38], and Szabo et al.,
unpublished data), the membrane-anchored serine protease
nevertheless is expressed in many burgeoning embryonic as well
as extraembryonic epithelia [39,40,41,42]. Furthermore, we have
previously shown that matriptase must be tightly regulated at the
post-translational level, for successful execution of several developmental
processes. Thus, loss of either of the two Kunitz-type
transmembrane serine protease inhibitors, hepatocyte growth
factor activator inhibitor (HAI)-1 or -2 or combined haploinsufficiency
for both inhibitors, is associated with uniform embryonic
lethality in mice [40,43]. Loss of HAI-1 or combined haploinsufficiency
for HAI-1 and HAI-2 causes mid-gestation embryonic
lethality due to failure to develop the placental labyrinth. Loss of
HAI-2, in turn, is associated with three distinct phenotypes: a)
Early embryonic lethality, b) mid-gestation lethality due to
placental labyrinth failure, and c) neural tube defects resulting in
exencephaly, spina bifida, and curly tail. All developmental defects
in HAI-1- and HAI-2-deficient embryos, however, are rescued in
whole or in part by simultaneous matriptase-deficiency, thus
demonstrating that a matriptase-dependent proteolytic pathway is
a critical morphogenic target for both protease inhibitors ([43,44],
this study).
In this study, we exploited the observation that HAI-1- and
HAI-2-deficient mice display matriptase-dependent embryonic
lethality with complete penetrance to perform a comprehensive
genetic epistasis analysis aimed at identifying additional components
of the matriptase proteolytic pathway. Specifically, we
generated mice with simultaneous ablation of either the Spint1
gene (encoding HAI-1) or the Spint2 gene (encoding HAI-2) along
with genes encoding candidate matriptase targets that are coexpressed
with the protease during development. We then
screened for the rescue of embryonic lethality or restoration of
HAI-1 and HAI-2-dependent morphogenic processes in these
double-deficient mice. This analysis identified prostasin as critical
to all matriptase-induced embryonic defects in both HAI-1- and
HAI-2-deficient mice. Paradoxically, however, although matriptase
autoactivates efficiently and prostasin is incapable of
undergoing autoactivation, we found that prostasin acts upstream
of matriptase in the developing embryo and is required for
conversion of the matriptase zymogen to active matriptase.
Finally, we explored the contribution of this newly identified
prostasin-matriptase pathway to protease-activated receptor
(PAR)-dependent signaling during neural tube formation [45]
and now provide evidence that the pathway may be separate from
the proteolytic machinery that mediates focal activation of PAR-2
during neural tube closure.
Results
Developmental defects in HAI-2–deficient mice tightly
correlate with matriptase expression levels
HAI-2-deficient (Spint2 2/2 ) mice were originally reported to
display embryonic lethality prior to embryonic day 8 (E8.0),
presenting with severe clefting of the embryonic ectoderm at E7.5
and a failure to progress to the headfold stage [44]. We previously
reported, however, that approximately 50% of HAI-2-deficient
mice complete early development but die at midgestation due to
defective placental branching morphogenesis [43]. However, the
genotyping strategy used in the latter study aimed at exploring the
contribution of matriptase to this embryonic demise and only
allowed for the discrimination of HAI-2-deficient mice on
matriptase-sufficient (wildtype, Spint2 2/2 ;St14 +/+ , or haploinsufficient,
Spint2 2/2 ;St14 +/2 ) backgrounds from a matriptase-deficient
(Spint2 2/2 ;St14 2/2 ) background. Therefore, to test the possibility
that early embryonic development of HAI-2-deficient mice is St14
gene dosage-dependent, we first analyzed the offspring of
interbred Spint2 +/2 ;St14 +/2 mice at various developmental stages.
This analysis revealed that the various developmental phenotypes
seen in HAI-2-deficient mice, indeed, were strongly dependent on
St14 gene dosage (Figure 1A). Thus, HAI-2-deficient embryos
carrying two wildtype matriptase alleles (St14 +/+ ), displayed early
lethality, as evidenced by only five percent of Spint2 2/2 ;St14 +/+
embryos developing beyond E9.0 and none past E10.5 (Figure 1A,
blue diamonds). Inactivation of one matriptase allele (Spint2 2/2 ;
St14 +/2 ), however, was sufficient to partially rescue this early
embryonic lethality of HAI-2-deficient mice (Figure 1A, red
squares). As reported previously [43], inactivation of both alleles of
matriptase (Spint2 2/2 ;St14 2/2 ) completely restored embryonic
survival and placental development and also reduced the
occurrence of neural tube defects associated with the loss of
HAI-2 (Figure 1A, green triangles and Table 1). Taken together,
these findings show that loss of HAI-2 may lead to three distinct
developmental phenotypes, dependent on the overall expression
level of matriptase (Table 1): (i) early embryonic lethality occurring
largely prior to E8.5, which can be partially rescued by matriptase
haploinsufficiency (Spint2 2/2 ;St14 +/2 ) and completely by matriptase
deficiency (Spint2 2/2 ;St14 2/2 ); (ii) placental defects resulting in
mid-gestation lethality, which are observed in Spint2 2/2 ;St14 +/2
embryos after E9.5, but are absent in Spint2 2/2 ;St14 2/2 embryos,
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Figure 1. Effect of St14 gene dosage and c-Met activity on embryonic development in HAI-1– and HAI-2–deficient mice. (A) Matriptase
haploinsufficiency partially restores early embryonic development of HAI-2 deficient mice. Relative frequency of Spint2 2/2 ;St14 +/+ (blue diamonds
and trend line), Spint2 2/2 ;St14 +/2 (red squares and trendline), and Spint2 2/2 ;St14 2/2 (green triangles and trendline) embryos in offspring from
interbred Spint2 +/2 ;St14 +/2 mice at E8.5–E15.5. The expected 25% Mendelian frequency is shown with the dotted trend line. 59–250 embryos were
genotyped at each stage. (B) Matriptase haploinsufficiency does not rescue development of HAI-1-deficient mice. Genotype distribution of E8.5–E11.5
embryos and newborn (P1) offspring from interbred Spint1 +/2 ;St14 +/2 mice. No living St14 +/+ ;Spint1 2/2 or St14 +/2 ;Spint1 2/2 embryos are observed
after E9.5. (C) Distribution of Spint1 genotypes in c-Met-expressing (Hgfr +/+ or Hgfr +/2 , blue bars) and c-Met-deficient (Hgfr 2/2 , green bars) embryos
from interbred Spint1 +/2 ;Hgfr +/2 mice at E11.5–13.5. Loss of c-Met activity does not improve embryonic survival of HAI-1-deficient mice. (D and E)
Distribution of Spint2 genotypes in c-Met-expressing (Hgfr +/+ or Hgfr +/2 , blue bars) and c-Met-deficient (Hgfr 2/2 , green bars) embryos from
Spint2 +/2 ;Hgfr +/2 6Spint2 +/2 ;Hgfr +/2 (D) or Spint2 +/2 ;Hgfr +/2 6Spint2 +/2 ;Hgfr +/2 ;St14 +/2 (E) breeding pairs at E9.5–10.5. Only St14 +/2 embryos are
shown in (E). Loss of c-Met does not improve survival of HAI-2-deficient embryos. (F) Frequency of exencephaly observed in 153 control
(Spint2 + ;Hgfr + ), 53 c-Met- (Spint2 + ;Hgfr 2/2 ), 24 HAI-2- (Spint2 2/2 ,Hgfr + ), and 6 c-Met and HAI-2 double- (Spint2 2/2 ;Hgfr 2/2 ) deficient embryos at E9.5.
Loss of c-Met activity fails to correct neural tube defects in HAI-2-deficient mice.
doi:10.1371/journal.pgen.1002937.g001
Table 1. Developmental defects observed in Spint1- and Spint-2-deficient mice as function of St14 expression.
Genotype Phenotype Penetrance
Spint1 2/2
Lack of placental labyrinth, embryonic lethality
at E10.5
St14 +/+ St14 +/2 St14 2/2
100% 100% (no rescue) 0% (complete rescue)
Spint2 2/2 Early embryonic lethality at E9.5 or earlier 100% 45% (partial rescue) 0% (complete rescue)
Incomplete differentiation of placental
N/A 100% (no rescue) 0% (complete rescue)
labyrinth, embryonic lethality at E10.5–E14.5
Neural tube defects
N/A
Exencephaly 95–100% 18% (partial rescue; P,0.0001)
Spina bifida 11% 13% (no rescue)
Curly tail 89% 62% (partial rescue?; not
significant)
doi:10.1371/journal.pgen.1002937.t001
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and (iii) neural tube defects observed at or after E8.5 in most
Spint2 2/2 ;St14 +/2 embryos, and partially rescued in Spint2 2/2 ;
St14 2/2 embryos and term offspring.
We next performed a similar analysis of the effect of St14 gene
dosage on the developmental defects and embryonic lethality
associated with HAI-1-deficiency by analyzing the offspring from
interbred Spint1 +/2 /St14 +/2 mice (Figure 1B). As shown previously
[40], a complete rescue of both the placental defects and
embryonic lethality was observed in HAI-1-deficient mice
expressing no matriptase (Spint1 2/2 ;St14 2/2 ). However, comparison
of HAI-1-deficient mice carrying one (Spint1 2/2 ;St14 +/2 )or
two (Spint1 2/2 ;St14 +/+ ) wildtype St14 alleles revealed identical
defects in placental labyrinth formation and mid-gestation
embryonic lethality occurring with complete penetrance
(Figure 1B, Table 1, and data not shown).
Activation of hepatocyte growth factor (HGF) does not
contribute to placental defects in HAI-1–deficient
embryos or early embryonic lethality and neural tube
defects in HAI-2–deficient mice
Matriptase is an efficient activator of proHGF [29,30] and
dysregulated matriptase activity recently was shown to promote
squamous cell carcinoma through activation of HGF-dependent c-
Met signaling [46]. Furthermore, both proHGF and its cognate
receptor c-Met are expressed during embryogenesis in both the
placenta and the embryo [47,48]. To investigate the involvement
of aberrant proHGF activation and c-Met signaling in the etiology
of the defects observed in HAI-1- and HAI-2-deficient embryos,
we took advantage of the fact that c-Met is only required for
embryonic development beyond E13.5 [47,48]. This enabled the
study of key HAI-1- and HAI-2-dependent morphogenic processes
in mice homozygous for a null mutation in Hgfr (Hgfr 2/2 ),
encoding c-Met. Analysis of embryos from interbred Spint1 +/2 ;
Hgfr +/2 mice at E11.5–E13.5 revealed only one surviving
Spint1 2/2 ;Hgfr 2/2 embryo, indicating that the loss of c-Met
activity does not restore placental development or embryonic
survival of HAI-1-deficient mice (Figure 1C, P,0.04, Chi-square
test, and data not shown). Likewise, no Spint2 2/2 ;Hgfr 2/2
embryos were detected beyond E9.5 (Figure 1D, P,0.02, Chisquare
test), indicating that the inactivation of c-Met signaling does
not prevent matriptase-induced early embryonic lethality in HAI-
2-deficient mice. Interbreeding Spint2 +/2 ;Hgfr +/2 ;St14 +/2 mice
allowed for the analysis of the impact of c-Met deficiency on the
formation of neural tube defects in HAI-2-deficient mice by
preventing early embryonic lethality (Figure 1E). However, all of
the Spint2 2/2 ;Hgfr 2/2 ;St14 +/2 embryos isolated at E9.5 from
these crosses presented with exencephaly (Figure 1F), suggesting
that c-Met signaling is not critically involved in the neural tube
defects caused by the absence of HAI-2. All Spint2 2/2 ;
Hgfr 2/2 ;St14 +/2 embryos displayed synthetic lethality after E9.5,
which precluded the direct analysis of the impact of c-Met loss on
the defects in placental differentiation caused by HAI-2 deficiency
(data not shown). Taken together, these findings suggest that
aberrant HGF-c-Met signaling does not contribute to the
matriptase-dependent defects in placentation in HAI-1-deficient
embryos, or early lethality and neural tube closure of HAI-2-
deficient embryos.
Reduced prostasin enzymatic activity prevents
developmental defects in both HAI-1– and HAI-2–
deficient mice
The GPI-anchored membrane serine protease, prostasin
(CAP1/PRSS8), is a well-validated downstream proteolytic target
for matriptase in the epidermis of mice and humans (see
Introduction). To explore the possibility that matriptase acts
through prostasin to cause the signature defects in embryonic
development of HAI-1- and HAI-2-deficient mice, we first
performed a detailed immunohistochemical analysis of prostasin
expression in the developing embryo by staining histological
sections from wildtype (Prss8 +/+ ) and littermate prostasin-deficient
(Prss8 2/2 ) embryos with prostasin antibodies. Interestingly,
prostasin was expressed in both the surface ectoderm, specifically
covering the converging neuroepithelium at the time of the neural
tube closure (Figure 2A and 2B, compare with 2C), and in the
developing placenta, where expression was detected as early as on
E8.5 and was present in the placental labyrinth in the entire period
of placental differentiation (Figure 2D, 2E, 2G, and 2H, compare
with 2F and 2I), thereby displaying co-expression with matriptase,
HAI-1 and HAI-2 [40,41,42,43,45]. We, therefore, next directly
determined the contribution of prostasin to the matriptasedependent
developmental defects of HAI-1- and HAI-2-deficient
mice. For this purpose, we exploited the fact that the spontaneous
mutant mouse strain, frizzy, recently was described to be
homozygous for a point mutation in the coding region of the
Prss8 gene (Prss8 fr/fr ). This mutation results in a non-conservative
V170D amino acid substitution in the prostasin protein [49].
Moreover, this mutant mouse strain completes development, but
displays an epidermal phenotype resembling mice carrying a
hypomorphic mutation in St14 [23], suggesting reduced expression
or enzymatic activity of V170D prostasin. Western blot and
immunohistochemical analysis of tissues from Prss8 fr/fr mice did
not reveal an obvious reduction in the level of V170D prostasin
expression when compared to wildtype prostasin in Prss8 +/+
littermates (data not shown). Therefore, to assess the enzymatic
activity of the mutant prostasin, we generated enteropeptidaseactivated
recombinant V170D prostasin, as well as enteropeptidase-activated
wildtype and catalytically inactive (S238A) prostasin
variants in HEK293T cells, as described previously [26]. These
recombinant proteins were released from the plasma membrane
by phosphatidylinositol-specific phospholipase C, activated with
enteropeptidase, and their enzymatic activity towards a prostasinselective
fluorogenic peptide substrate (Figure 2J) as well as their
ability to form enzymatic activity-dependent covalent complexes
with the serpin, protease nexin-1 (PN-1) (Figure 2K), were tested.
As expected, wildtype recombinant prostasin exhibited easily
detectable hydrolytic activity towards the fluorogenic peptide
(Figure 2J, red line) and formed SDS-stable complexes with PN-1
(Figure 2K, compare lanes 3 and 4), while prostasin not activated
by enteropeptidase and the catalytically inactive S238A mutant
and exhibited no detectable hydrolytic activity (Figure 2J, black
and grey lines) or PN-1 binding (Figure 2K and Figure S1A, lanes
2 and 12). V170D prostasin displayed a low residual enzymatic
activity that was above the baseline level, as defined by the
catalytically inactive S238A variant, and corresponded to about
6% of the activity of wildtype prostasin (Figure 2J, blue line), while
complex formation with PN-1 could not be detected (Figure 2K
and Figure S1A, compare lanes 7 and 8). Taken together, these
data indicated that V170D prostasin, expressed by the Prss8 fr
allele, displays greatly reduced enzymatic activity. We, therefore,
next interbred Spint1 +/2 ;Prss8 fr/+ and Spint1 +/2 ;Prss8 fr/fr mice and
analyzed the distribution of Spint1 alleles in the newborn offspring
from these crosses. Consistent with our previous findings, loss of
HAI-1 was not compatible with embryonic survival of mice
carrying a wildtype prostasin allele (Spint1 2/2 ;Prss8 fr/+ ) (Figure 3A,
blue bars). Interestingly, however, HAI-1-deficient mice carrying
two mutant prostasin alleles (Spint1 2/2 ;Prss8 fr/fr ) developed to term
(Figure 3A, green bars), although they were found at a frequency
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Figure 2. Prostasin expression in embryonic and extraembryonic tissues. (A–C) Immunohistochemical detection of prostasin at E8.5 in
epithelial cells of surface ectoderm (examples with arrows in A and B) overlying the cranial neural tube region. Specificity of staining is shown by the
absence of staining of Prss8 2/2 surface ectoderm (arrow in C). Filled arrowhead shows non-specific staining of yolk sac. No expression was observed
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Embryonic Cell Surface Serine Protease Cascade
in the neuroepithelium (A and B, open arrowheads). (D–F) Immunohistochemical detection of prostasin in the chorionic ectoderm (examples with
arrows) of mouse placenta at E8.5. Specificity of staining is shown by the absence of staining of Prss8 2/2 chorionic ectoderm (F). Filled arrowheads in
D and F shows non-specific staining of trophoblast giant cells. No expression was detected in the trophoblast stem cell-containing chorionic
epithelium (open arrowhead in E). (G–I) Immunohistochemical detection of prostasin in the placental labyrinth (examples with arrows in G and H) of
mouse placenta at E12.5. Specificity of staining is shown by the absence of staining of the Prss8 2/2 labyrinth (I). No expression was detected in the
trophoblast stem cell-containing chorionic epithelium (open arrowhead in H). Scale bars: A, C, D, F, G, and I, 100 mm; B, E, and H, 25 mm. (J) Enzymatic
activity of wildtype (red), V170D (blue), S238A (grey), and zymogen (black) forms of prostasin. Prostasin variants were incubated with 50 mM pERTKR-
AMC fluorogenic peptide at 37uC. V170D prostasin exhibited about 6% of the amidolytic activity of wildtype prostasin. No activity of catalytically
inactive prostasin or prostasin zymogen was detected. (K) Western blot detection of SDS-stable complexes between prostasin and protein nexin-1
(PN-1). Wildtype zymogen (lanes 1 and 2), activated wildtype (lanes 3 and 4), V170D (frizzy) zymogen (lanes 5 and 6), activated V170D (lanes 7 and 8),
S238A zymogen (lanes 9 and 10), and activated S238A (lanes 11 and 12) prostasin variants were incubated with (lanes 2, 4, 6, 8, 10, and 12) or without
(lanes 1, 3, 5, 7, 9, and 11) 250 ng of recombinant human PN-1. Wildtype, but not V170D or S238A variants of prostasin formed SDS-stable complexes
with PN-1. Positions of pro-prostasin, activated prostasin (migrating slightly faster than the zymogen due to removal of the 12 aa propeptide that is
not detected after 4–12% SDS/PAGE with anti-prostasin antibody), and prostasin/PN-1 complexes are indicated. Positions of molecular weight
markers (kDa) are shown on left.
doi:10.1371/journal.pgen.1002937.g002
that was slightly lower than the expected Mendelian distribution
(20/127, 16% vs. expected 31.75/127, 25%, P,0.05, Chi-square
test). Furthermore, morphometric analysis showed that reduced
prostasin activity fully restored placental labyrinth formation in
HAI-1-deficient embryos, as evidenced by normal histological
appearance of the labyrinth (Figure 3B–3G), thickness of the
labyrinth layer (Figure 3H) and labyrinth vessel density (Figure 3I)
of Spint1 2/2 ;Prss8 fr/fr embryos. Furthermore, macroscopic and
histological analysis of embryos extracted between E11.5 and
E13.5 failed to reveal any obvious developmental abnormalities
within either embryonic or extraembryonic tissues of
Spint1 2/2 ;Prss8 fr/fr mice (data not shown), and these mice were
outwardly indistinguishable from their Prss8 fr/fr littermates at
weaning and when followed for up to one year (Figure 3J). Taken
together, these data show that the matriptase-mediated developmental
defects in HAI-1-deficient mice are prostasin-dependent.
To determine the impact of diminished prostasin activity on the
developmental defects associated with HAI-2-deficiency, we next
analyzed neural tube closure, placental differentiation, and overall
survival of the offspring of interbred Spint2 +/2 ;Prss8 fr/+ mice.
Analysis of the genotype distribution of embryos at E9.5–11.5 did
not identify any HAI-2-deficient embryos carrying at least one
wildtype Prss8 allele (Spint2 2/2 ;Prss8 +/+ or Spint2 2/2 ;Prss8 fr/+ )
(Figure 4A). Interestingly, however, HAI-2-deficient embryos
carrying two mutant Prss8 alleles (Spint2 2/2 ;Prss8 2fr/fr ) were found
in the expected Mendelian ratio as late as E13.5–15.5 (Figure 4B,
green bars). Furthermore, genotyping of newborn offspring
revealed the presence of living Spint2 2/2 ;Prss8 fr/fr pups
(Figure 4C, green bars), although they were found at slightly
lower than expected frequency (15% vs. expected 25%, P,0.06,
Chi-square test). These data strongly suggest that matriptase and
prostasin act as part of a single proteolytic cascade to cause
developmental defects in HAI-2-deficient mice. If this were the
case, we hypothesized that lowering the activity of this cascade
even further by eliminating one St14 allele from Spint2 2/2 ;Prss8 fr/fr
embryos should additionally improve the term survival of HAI-2-
deficient mice. Indeed, genotyping of born offspring from
interbred Spint2 +/2 ;Prss8 fr/+ ;St14 +/2 mice showed a normal
distribution of Spint2 alleles in Prss8 fr/fr ;St14 +/2 pups (Figure 4C,
red bars), further suggesting that failure to regulate a proteolytic
pathway including matriptase and prostasin accounts for all of the
embryonic lethality caused by loss of HAI-2.
As reported previously, neural tube defects, including exencephaly,
spina bifida, and curly tail were seen in 95–100% of
Spint2 2/2 ;Prss8 fr/+ ;St14 +/2 mice ([43], this study). Examination of
embryonic and extraembryonic tissues from Spint2 2/2 ;Prss8 fr/fr
and Spint2 2/2 ;Prss8 fr/fr ;St14 +/2 embryos, however, revealed that
reduced prostasin activity sufficed to almost completely rescue the
defects in both neural tube closure and placental differentiation
caused by HAI-2 deficiency. Thus, macroscopic (Figure 4D and
4E) and histological (Figure 4F and 4G) examination of
Spint2 2/2 ;Prss8 fr/fr ;St14 +/+ and Spint2 2/2 ;Prss8 fr/fr ;St14 +/2 embryos
showed that, respectively, 5% and 0%, of these embryos
exhibited exencephaly when analyzed after E9.5 (Figure 4H), and
no embryos with either spina bifida or curly tail were observed
(data not shown). Similarly, histological analysis of placental tissues
from E10.5–E13.5 Spint2 2/2 ;Prss8 fr/fr or Spint2 2/2 ;Prss8 fr/fr ;St14 +/
2 embryos did not reveal any of the stereotypic defects associated
with HAI-2 deficiency (Figure 4I and 4J). Thus, the overall
appearance of the placental layers (Figure 4I and 4J), the thickness
of the placental labyrinth (Figure 4K), and the number of fetal
vessels within the labyrinth (Figure 4L), all were comparable to the
HAI-2-sufficient littermate controls. In conclusion, these data
document an essential role of prostasin in the etiology of all of the
developmental defects previously observed in HAI-2-deficient
mice.
Prostasin is required for the activation of matriptase
during development
Matriptase was previously identified as an essential proteolytic
activator of prostasin in the epidermis, and the near ubiquitous colocalization
of the two membrane serine proteases in the epithelial
compartment of most other adult tissues indicate that this
matriptase-prostasin proteolytic pathway may be operating in
multiple epithelia to maintain tissue homeostasis [23,24,25,26].
The genetic epistasis analysis performed above provided strong
evidence that matriptase and prostasin also are part of a single
proteolytic cascade in the context of embryonic development.
Furthermore, the striking overlap in expression of the two
proteases documented earlier in the surface ectoderm during
neural tube closure (see above) was also observed in the developing
placenta (compare Figure 5A and 5B). To further investigate the
functional interrelationship between the two proteases, we
analyzed the levels of the activated forms of matriptase and
prostasin in embryonic and placental tissues from matriptase-
(St14 2/2 ) or prostasin- (Prss8 2/2 ) deficient mice at E11.5.
Trypsin-like serine proteases are activated by autocatalytic or
heterocatalytic cleavage after an arginine or lysine residue, located
in a conserved activation motif within the catalytic domain.
Activation cleavage severs the bond between the catalytic domain
and upstream accessory domains, but the activated protease
domain remains connected to upstream accessory domains by a
disulfide bond [50]. Zymogen activation, therefore, can be
detected by a mobility shift in reducing SDS-PAGE gels, which
breaks the disulfide bond that keeps the two domains together.
Direct detection of active matriptase in placental tissues by western
blot, however, proved unsuccessful due to low signal intensity and
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Figure 3. Reduced prostasin activity restores placental development and embryonic survival of HAI-1–deficient mice. (A) Distribution
of genotypes of born offspring of intercrossed Spint1 +/2 ;Prss8 fr/2 mice. No Spint1 2/2 mice expressing one or two wildtype prostasin alleles (Prss8 +/+ or
Prss8 +/fr , blue bars) were identified, while Spint1 2/2 embryos carrying two mutant prostasin alleles (Prss8 fr/fr , green bars) were found in near-expected
frequency. (B–G) Representative low (B–D) and high (E–G) magnification images showing the histological appearance of H&E-stained placental tissues
of (Spint1 + ;Prss8 + ) (B and E), (Spint1 2/2 ;Prss8 + ) (C and F), and (Spint1 2/2 ;Prss8 fr/fr ) (D and G) embryos at E11.5. The thickness of the placental labyrinth
(two-sided arrows between the dotted lines in B–D), as well as the number of fetal vessels (E–G, arrows) and lacunae filled with maternal blood (E–G,
arrowheads) within the labyrinth is markedly reduced in prostasin-sufficient (C and F), but not in prostasin-deficient (D and G) Spint1 2/2 embryos,
when compared to the controls (B and E). (H, I) Quantification of the maximum thickness of the labyrinth layer (H) and the number of fetal vessels in
the placental labyrinth (I) of Spint1 + ;Prss8 + , Spint1 + ;Prss8 fr/fr , Spint1 2/2 ;Psrr8 + , and Spint1 2/2 ;Psrr8 fr/fr embryos at E11.5. The thickness of the labyrinth
and fetal vessel density were strongly diminished in HAI-1-deficient mice but completely restored in HAI-1-deficient mice with low prostasin activity.
(J) Outward appearance of one-year-old Spint1 2/2 ;Prss8 fr/fr and littermate Spint1 + ;Prss8 + mice. ***, p,0.0001, Student’s t-Test, two tailed. Scale bars:
B–D, 100 mm; E–G, 25 mm.
doi:10.1371/journal.pgen.1002937.g003
a strong cross reactivity of available anti-matriptase antibodies
with unrelated antigens. Similarly, direct detection of active
prostasin by western blot failed due to the small difference in the
electrophoretic mobility of the zymogen and the active form of the
enzyme (data not shown). In order to circumvent these problems,
we instead determined the amount of active matriptase and
prostasin that formed inhibitor complexes with endogenous HAI-1
in embryonic tissues from wildtype, matriptase-, and prostasindeficient
embryos. Immunoprecipitation of protein extracts using
anti-mouse HAI-1 antibodies followed by western blot with
prostasin antibodies detected the presence of the 38 kDa band
in the placentas of wildtype mice (Figure 5C, lanes 2 and 4) and
matriptase-deficient mice (Figure 5C, lane 3). This band was not
detected in placental extracts from prostasin-deficient embryos
(Figure 5C, lane 1) or when anti-HAI-1 antibodies were omitted
from the assay (Figure 5D, compare lanes 1 and 2), indicating that
it represents the active form of prostasin released from an
inhibitory complex with HAI-1. In support of this, when prostasin
from either matriptase-deficient or littermate wildtype control
placental tissues was released from the immunoprecipitated HAI-
1-prostasin complexes by brief exposure to low pH, it was able to
form SDS-stable complex with PN-1, which requires the catalytic
activity of prostasin (Figure 5C, compare lane 3 with 5 and lane 4
with 6). Quantification of the amount of active prostasin in
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Figure 4. Reduced prostasin activity restores placental differentiation, embryonic survival, and neural tube closure in HAI-2–
deficient mice. (A) Distribution of Spint2 genotypes in prostasin-sufficient (Prss8 +/+ or Prss8 +/fr ) E9.5–11.5 offspring from interbred Spint2 +/2 ;Prss8 fr/+
mice. No Spint2 2/2 embryos were observed (P,0.025, Chi-square test). (B) Distribution of Spint2 genotypes in prostasin-sufficient (Prss8 +/+ or Prss8 +/fr ,
blue bars) and prostasin-deficient (Prss8 fr/fr , green bars) mouse embryos from interbred Spint2 +/2 ;Prss8 fr/+ mice at E13.5–15.5. No prostasin-expressing
Spint2 2/2 embryos were observed (P,0.001, Chi-square test), while survival of prostasin-deficient Spint2 2/2 embryos was restored. (C) Distribution of
Spint2 genotypes in newborn prostasin-sufficient, matriptase wildtype (Prss8 +/+ or Prss8 +/fr ;St14 +/+ , blue bars), prostasin-deficient, matriptase wildtype
(Prss8 fr/fr ;St14 +/+ , green bars), and prostasin-deficient, matriptase haploinsufficient (Prss8 fr/fr ;St14 +/2 , red bars) offspring from Spint2 +/2 ;Prss8 fr/2
6Spint2 +/2 ;Prss8 fr/+ ;St14 +/2 breeding pairs. Reduced prostasin activity restored embryonic survival of Spint2 2/2 mice partially in matriptase wildtype
and completely in matriptase haploinsufficient mice. (D–G) Macroscopic (D and E) and histological (H&E staining) (F and G) appearance of the HAI-2-
deficient, matriptase- and prostasin-sufficient (Spint2 2/2 ;Prss8 +/+ or Prss8 +/fr , St14 +/2 , D and F) or HAI-2- and prostasin-deficient, matriptase-sufficient
(Spint2 2/2 ;Prss8 fr/fr , St14 +/+ or St14 +/2 ) (E and G) embryos at E9.5. HAI-2 deficiency prevents convergence of neural folds in the cranial region of neural
tube (D and F, arrows) leading to exencephaly. Convergence and fusion of neural folds are restored in HAI-2-deficient mice with low prostasin activity
(E and G, arrows). Presence of medial (F, open arrowhead) and absence of dorsolateral (F, arrowheads) hinge points. (H) Frequency of exencephaly in
E9.5–18.5 Spint2 2/2 embryos with different levels of prostasin activity (Prss8 +/+ , Prss8 fr/+ or Psrr8 fr/fr ) and matriptase (St14 +/+ , St14 +/2 or St14 2/2 ). The
frequency of neural tube defects is inversely correlated with the combined number of wildtype Prss8 and St14 alleles. A total of 524 embryos were
analyzed. (I–L) Histological appearance (H&E staining) (I and J), thickness of placental labyrinth (K), and number of fetal vessels within the labyrinth (L)
in the placentas of HAI-2 and prostasin-sufficient (Spint2 + ;Prss8 + ) and HAI-2 and prostasin double-deficient (Spint2 2/2 ;Prss8 fr/fr ) embryos at E12.5.
Reduced prostasin activity restores differentiation of placental labyrinth in Spint2 2/2 mice to levels not significantly (N.S.) different from wildtype
littermate controls. Arrows in I and J show examples of fetal vessels. Scale bars: F, 50 mm G, I, and J, 100 mm.
doi:10.1371/journal.pgen.1002937.g004
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Figure 5. Prostasin is required for the activation of matriptase during placental differentiation. (A and B) Expression of prostasin (A) and
matriptase (B) in placental tissues of wildtype mice at E11.5. Both proteins were expressed in the chorionic (arrows) and labyrinthine (arrowheads)
trophoblasts. (C) Western blot detection of active prostasin in the fetal part of the placenta of wildtype (Prss8 +/+ and St14 +/+ , lanes 2, 4, and 6),
prostasin-deficient (St14 +/+ ;Prss8 2/2 ) (lane 1), and matriptase-deficient (St14 2/2 ;Prss8 +/+ ) (lanes 3 and 5) embryos at E11.5 after immunoprecipitation
with anti-mouse HAI-1 antibodies. Immunoprecipitated proteins in lanes 5 and 6 were acid-exposed to dissociate prostasin-HAI-1 complexes, and
then incubated with PN-1 prior to western blot analysis. Positions of bands corresponding to active prostasin, prostasin/PN-1 complex, as well as nonspecific
signals of IgG heavy and light chains are indicated on the right. Positions of molecular weight markers (kDa) are shown on the left. (D)
Omission of anti-HAI-1 antibody resulted in loss of detectable prostasin (compare lanes 1 and 2), indicating that the detected prostasin formed
complexes with HAI-1. (E) Quantification of the relative amount of active prostasin in wildtype and matriptase placentae by densitometric scanning of
prostasin western blots of HAI-1 immunoprecipitated material from (Prss8 2/2 ;St14 +/+ ,N=3,Prss8 +/+ ;St14 2/2 , N = 3, and Prss8 +/+ ;St14 +/+ , N = 6). Data
are shown as mean 6 standard deviation (N.S., not significant). (F and G) Western blot detection of active matriptase in the fetal part of the placenta
at E11.5 (F) after anti-HAI-1 immunoprecipitation, and in the epidermis of newborn skin (G) of wildtype (Prss8 +/+ and St14 +/+ ) (F, lanes 1, and 3, and G,
lane 2), prostasin-deficient (St14 +/+ ;Prss8 2/2 ) (F, lane 4 and G, lane 1), and matriptase-deficient (St14 2/2 ;Prss8 +/+ ) (F, lane 2, and G, lane 3) embryos. A
30 kDa band representing the active serine protease domain of matriptase (Mat SPD) was present in extracts from wildtype (lanes 1 and 3 in F), but
not in matriptase- (lane 2 in F) or prostasin-deficient (lane 4 in F) placenta. Zymogen (Mat FL) and active (Mat SPD) forms of matriptase were detected
in extracts from both wildtype and prostasin-deficient, but not matriptase-deficient epidermis. (H–H0) Immunohistochemical staining of matriptase in
control Prss8 + (H) and prostasin-deficient Prss8 2/2 (H9) placenta at E11.5. Specificity of staining of chorionic and labyrinthine trophoblasts (examples
with arrows) is shown by the absence of staining of corresponding cells in St14 2/2 placenta (H0). Insets in H and H9 are parallel sections stained with
prostasin antibodies. Open arrowheads in H–H0 show examples of non-specific staining. Scale bars: A, B, H, H9, and H0, 50mm.
doi:10.1371/journal.pgen.1002937.g005
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wildtype and prostasin-deficient placentae by densitometric scans
of western blots showed that the loss of matriptase did not affect
the amount of active prostasin (Figure 5E). Taken together, these
data suggest that the developing placenta does not require
matriptase for the activation of prostasin.
Detection of matriptase by western blot after immunoprecipitation
with anti-HAI-1 antibodies revealed the presence of a
30 kDa band corresponding to the activated matriptase serine
protease domain in wildtype placental tissues, but not matriptasedeficient
placental tissues (Figure 5F, compare lanes 1 and 2).
Surprisingly, however, the active form of matriptase was also
absent in the extracts from prostasin-deficient placentae (Figure 5F,
compare lanes 3 and 4). The absence of matriptase was observed
in four independent experiments using placentae from a total of
seven prostasin-deficient mice and their prostasin-sufficient littermate
controls (Figure S1B and data not shown). As expected,
analysis of skin extracts from prostasin-deficient newborn mice and
wildtype littermate controls using the same western blot conditions
clearly showed the presence of the active form of matriptase in
both the control and prostasin-deficient mice (Figure 5G, compare
lanes 1 and 2 with lane 3), demonstrating that differences in the
functional relationship between the two proteases exist in different
tissues. Immunohistochemistry of placentae from littermate
control and prostasin-deficient embryos showed no obvious
difference in levels or pattern of matriptase expression (compare
Figure 5H and 5H9).
To further substantiate the above findings, we next determined
if prostasin could serve as an activator of matriptase in a
reconstituted cell-based assay. For this purpose, we transiently
transfected HEK-293 cells with expression vectors encoding HAI-
1 (to allow for efficient matriptase expression) and wildtype or
catalytically inactive matriptase. The transfected cells were then
exposed to soluble recombinant prostasin or vehicle, and
matriptase activation was analyzed six hours later by western blot
of cell lysates (Figure 6A) or conditioned medium (Figure 6B).
Interestingly, soluble prostasin efficiently activated matriptase, as
evidenced by the large increase in the amount of the liberated
matriptase serine protease domain (Mat SPD, Figure 6A and 6B,
compare lanes 1 and 2) after reducing SDS/PAGE, and a
corresponding diminution of the amount of matriptase zymogen
(Mat SEA, Figure 6A and 6B, compare lanes 1 and 2). Activation
site cleavage of matriptase by prostasin did not require matriptase
catalytic activity, as shown by the increased amount of the isolated
matriptase serine protease domain in prostasin-treated cells
expressing a catalytically inactive matriptase (Figure 6A and 6B,
compare lanes 3 and 4). Similar results were obtained when
matriptase-transfected HEK-293 cells were transfected with a
prostasin expression vector, rather than being treated with soluble
prostasin (data not shown). To investigate if the prostasin-activated
matriptase displayed functional activity, the HEK-293 cells
described above were also transfected with a PAR-2 expression
vector and a serum response element (SRE)-luciferase reporter
plasmid to measure PAR-2 activity (Figure 6C). Exposure of
serum-starved cells to soluble prostasin resulted in a large increase
in luciferase activity in cells transfected with wildtype matriptase
(Figure 6C, left panels), but not in cells transfected with
catalytically inactive matriptase (Figure 6C, second panels from
left), with HAI-1 alone (Figure 6C, second panels from right) or
with empty vector (Figure 6C, right panels). Taken together, the
data indicate that prostasin can proteolytically activate matriptase
and is critical for the generation of active matriptase during
placental development. Detection of active matriptase and
prostasin in the embryo by western blot or by anti-HAI-1
immunoprecipitation failed to detect either of the proteases, likely
due to the restricted expression of both proteins (data not shown).
Developmental defects in HAI-2–deficient embryos are
not caused by aberrant activity of the epithelial sodium
channel
Both matriptase and prostasin have been reported to activate
the epithelial sodium channel (ENaC) in cell-based assays, and
prostasin is a critical regulator of ENaC activity during alveolar
fluid clearance in mouse lungs and likely regulates ENaC activity
in many other adult organs [51,52]. Immunohistological analysis
of embryonic tissues at E11.5 revealed strong expression of ENaC
in the developing labyrinth layer of the placenta (Figure 6D). No
ENaC expression was detected in the embryo proper (data not
shown). To investigate a possible involvement of ENaC in the
etiology of prostasin-matriptase-induced developmental defects in
HAI-2-deficient mice, pregnant females from Spint2 +/2 mice bred
to Spint2 +/2 ;St14 +/2 mice were treated daily between E5.5–8.5
with the pharmacological inhibitor of ENaC activity, amiloride,
which is known to cross the feto-maternal barrier [53]. Genotyping
of embryos extracted at E9.5 from these crosses did not
identify any Spint2 2/2 ;St14 +/+ embryos (Figure S1C), indicating
that the inhibition of ENaC activity is not sufficient to prevent
early embryonic lethality resulting from the loss of HAI-2.
Furthermore, all of the seven Spint2 2/2 ;St14 +/2 embryos identified
in this experiment exhibited exencephaly, suggesting that
ENaC activity is not critically involved in the etiology of neural
tube defects in HAI-2-deficient mice (Figure 6E). Similarly, genetic
inactivation of the a subunit of ENaC (encoded by Scnn1a), which
is necessary for channel activity in vivo [54,55], failed to rescue
embryonic development of HAI-2-deficient animals, as evidenced
by a complete absence of any surviving Spint2 2/2 ;Scnn1a 2/2
double-deficient embryos at E9.5 from Spint2 +/2 ;Scnn1a +/2 mice
bred to Spint2 +/2 ;Scnn1a +/2 mice (Figure 6F) and the failure of
Spint2 2/2 ;Scnn1a 2/2 double-deficient mice to appear in the
newborn offspring from Spint2 +/2 ;Scnn1a +/2 mice bred to
Spint2 +/2 ;Scnn1a +/2 ;St14 +/2 mice (Figure 6G). Taken together,
these data do not support the critical involvement of aberrant
ENaC activity in the developmental defects resulting from lack of
HAI-2 regulation of the prostasin-matriptase proteolytic pathway.
Excess PAR-2 signaling does not cause developmental
defects in HAI-2–deficient mice
Matriptase and prostasin are co-expressed with PAR-2 in
surface ectoderm during neural tube closure ([43,45], this study),
and matriptase displays extraordinarily favorable activation
kinetics towards PAR-2 in cell-based assays [43,45]. Furthermore,
activation of PAR-2 (encoded by the F2rl1 gene) was recently
shown to contribute to neural tube closure (see below). These data
suggested that some, or all, of the prostasin- and matriptasedependent
defects in HAI-2-deficient mice could be caused by
excess PAR-2 signaling. To test this hypothesis, we interbred
Spint2 +/2 ;F2rl1 +/2 mice and genotyped the ensuing embryos at
E9.5. This analysis failed to identify any Spint2 2/2 ;F2rl1 2/2
embryos (Figure 7A). Thus, the loss of PAR-2 activity is not
sufficient to overcome matriptase- and prostasin-dependent early
embryonic lethality in HAI-2-deficient mice. When the early
embryonic survival was improved by matriptase haploinsufficiency
(see above), analysis of neural tubes at E9.5 revealed exencephaly
in 100% of Spint2 2/2 ;F2rl1 2/2 St14 +/2 embryos, identical to the
frequency of defects observed in littermate HAI-2-deficient
embryos expressing PAR-2 (Spint2 2/2 ;F2rl1 +/+ or F2rl1 +/2 ;
St14 +/2 ) (Figure 7B). Thus, excess PAR-2 activation does not
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Embryonic Cell Surface Serine Protease Cascade
Figure 6. Prostasin activates matriptase on the surface of HEK293 cells. (A and B) Western blot detection of matriptase in cell lysates (A) and
in the conditioned medium (B) from HEK293 cells transiently transfected with wildtype recombinant human matriptase and HAI-1 expression vectors
(lanes 1 and 2), catalytically inactive (S805A) matriptase and HAI-1 (lanes 3 and 4), HAI-1 alone (lanes 5 and 6), and cells transfected with a control
empty vector (lanes 7 and 8) that were incubated with (lanes 2, 4, 6, and 8) or without (lanes 1, 3, 5, and 7) 100 nM soluble recombinant human
prostasin. Addition of prostasin promoted conversion of the matriptase zymogen to its activated two-chain form. Positions of bands corresponding
to full length matriptase (Mat FL), matriptase pro-enzyme processed by autocatalytic cleavage within the SEA domain (Mat SEA), and activated
matriptase serine protease domain (Mat SPD) are indicated on the right. Positions of molecular weight markers (kDa) are shown on left. (C)
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Embryonic Cell Surface Serine Protease Cascade
Quantification of the activation of PAR-2 in HEK293 cells expressing recombinant human PAR-2 in combination with wildtype (WT) or inactive (S805A)
variants of matriptase and HAI-1, HAI-1 alone, or transfected with an empty vector, incubated without (blue bars) or with (red bars) 100 nM soluble
recombinant human prostasin. Prostasin induced matriptase activity-dependent activation of PAR-2. (D) Immunohistochemical analysis of the
expression of the gamma subunit of the epithelial sodium channel (ENaC) in placenta of control mice at E11.5. The expression was detected in the
populations of chorionic (arrow) and labyrinthine (arrowhead) trophoblasts. Scale bar: 50 mm. (E) Frequency of exencephaly in amiloride-treated
wildtype (Spint2 +/+ , N = 56), untreated HAI-2-deficient (Spint2 2/2 , N = 12) and amiloride-treated HAI-2-deficient (Spint2 2/2 , N = 7) embryos at E9.5.
Amiloride treatment failed to rescue neural tube defects in Spint2 2/2 ; St14 +/2 embryos. (F) Distribution of Spint2 genotypes in ENaC-expressing
(Scnn1a +/+ or Scnn1a +/2 , blue bars) and ENaC-deficient (Scnn1a 2/2 , green bars) offspring from Spint2 +/2 ;Scnn1a +/2 6Spint2 +/2 ;Scnn1a +/2 breeding
pairs at E9.5. Loss of ENaC expression did not rescue early embryonic lethality in Spint2 2/2 mice. (G) Distribution of Spint2 genotypes in matriptasehaploinsufficient
ENaC-expressing (St14 +/2 ;Scnn1a +/+ or Scnn1a +/2 , blue bars) and ENaC-deficient (St14 +/2 ; Scnn1a 2/2 , green bars) offspring from
Spint2 +/2 ;Scnn1a +/2 , St14 +/2 6Spint2 +/2 ;Scnn1a +/2 ;St14 +/+ breeding pairs at birth. Loss of ENaC expression did not rescue overall embryonic survival in
Spint2 2/2 ; St14 +/2 mice.
doi:10.1371/journal.pgen.1002937.g006
appear to be critically involved in the etiology of neural tube
defects in HAI-2-deficient mice.
Loss of matriptase does not cause neural tube defects in
PAR-1–deficient embryos
Rac1 activation in surface ectoderm through G i , initiated by
either PAR-1 or PAR-2 activation was recently shown to be
required for neural tube closure. Thus, mice with combined, but
not single, deficiency in PAR-1 and PAR-2 display exencephaly
with high frequency [45]. As matriptase and prostasin are coexpressed
with PAR-2 in surface ectoderm during neural tube
closure ([43,45], this study), we next investigated if the prostasinmatriptase
cascade identified in the current study contributes to
physiological PAR-2 activation during neural tube closure. To test
this, we generated mice with combined deficiency in PAR-1
(encoded by the F2r gene) and matriptase (F2r 2/2 ;St14 2/2 ). If
matriptase was essential for the activation of PAR-2 during neural
tube closure, these mice should phenocopy mice with a combined
PAR-1 and PAR-2 deficiency, including embryonic lethality and
high susceptibility to cranial neural tube defects [45]. However,
analysis of the midgestation embryos from intercrossed
F2r +/2 ;St14 +/2 mice yielded the expected distribution of all
genotypes (Figure 7C). Furthermore, none of 20 observed F2r 2/
2 ;St14 2/2 mice displayed cranial neural tube defects, although the
PAR-1 deficiency alone or in combination with matriptase
deficiency occasionally led to defects in the closure of the posterior
neural tube, resulting in spina bifida and curly tail (Figure 7D).
HAI-2 and PAR-1/PAR-2 regulate different stages of
neural tube development
The lack of functional interaction between prostasin-matriptase
and PAR-1/PAR-2 regulated signaling pathways evidenced from
the above experiments suggested that the two pathways are either
involved in two essential, non-redundant mechanisms regulating
the same steps of neural tube closure, or that they may regulate
different stages of the process. To distinguish between the two
possibilities, we performed a detailed morphologic comparison of
the neural tube defects caused by loss of HAI-2 and by the
combined loss of PAR-1 and PAR-2. Macroscopic analysis showed
significant differences in the types of neural tube defects in the two
mutant mouse strains. In PAR-1 and PAR-2 double-deficient
mice, the defects were almost exclusively restricted to the
hindbrain region of the cranial neural tube (Figure 7E and
Table 2). In addition, PAR-1 and PAR-2 double-deficient embryos
did not exhibit any obvious abnormalities during the early stages
of neural tube closure, as all of the embryos analyzed before E10.5
showed normal elevation and conversion of the opposing neural
folds, as well as the completion of the neural fold fusion at initial
closure points 1 and 2 at hindbrain/cervix and forebrain/
midbrain boundaries, respectively (compare Figure 7F with 7G,
and 7F9 with 7G9). As a result, the neural tube defects in these
mice were generally only obvious after E10.5 (compare Figure 7H
with 7I and 7J), and were generally restricted to the hindbrain
region of the neural tube, with less than five percent exhibiting
exencephaly that extended to the midbrain region (Figure 7I and
Table 2). In contrast, HAI-2 deficiency was generally associated
with a failure of cranial neural tube closure that was obvious at
E9.5 or earlier, and was due to the inability of neural folds to
elevate properly, and to come into juxtaposition necessary for the
fusion (compare Figure 7F with 7K, and 7F9 with 7K9). The fusion
at closure point 1 of HAI-2-deficient mice was completed in all
embryos analyzed at E9.5 or later, and no case of craniorachischisis
was observed (Table 2). However, 10 percent of Spint2 2/2
embryos failed to initiate fusion at closure point 2, resulting in
exencephaly that extended from forebrain region to the hindbraincervical
boundary (Figure 7K and 7K9). In addition, even in the
embryos that successfully initiated the fusion at closure point 2, the
exencephaly was more extensive than the one observed in PAR-1
and PAR-2 double-deficient embryos, typically spanning the entire
midbrain and hindbrain regions (compare Figure 7L and 7I).
Finally, histological analysis of affected embryos at E9.5 showed
that dorsolateral hinge points (DLHPs) critical for the final stages
of the neural tube closure were absent in 97 percent of Spint2 2/2
embryos (Figure 4D and 4F, Figure 7K and 7K9, and Table 2),
while F2r 2/2 ;F2rl1 2/2 embryos generally exhibited DLHP
formation indistinguishable from wildtype littermate controls
(Figure 7G, 7G9, and 7M, compare to Figure 4E, 4G, Figure 7F
and 7F9). Thus, substantial differences are observed in the
location, frequency, extent, and onset of the neural tube defects
of HAI-2-deficient mice and PAR-1 and PAR-2 double-deficient
mice, further indicating the independent roles of, respectively,
repression and activation of the two protease-regulated pathways
in distinct stages of neural tube formation.
Discussion
In this study, we exploited the uniform matriptase-dependent
embryonic lethality of mice deficient in hepatocyte growth factor
activator inhibitors as a means to genetically identify novel
molecules and pathways regulating and being regulated by
matriptase in the developing embryo by epistasis analysis. This
analysis resulted in a number of unexpected findings. First, we
found that prostasin is an essential component of the matriptasedependent
molecular machinery that causes early embryonic
lethality, derails placental labyrinth formation, and causes defects
in neural tube closure in these mice. This shows that both proteins
are expressed, are active, functionally interact, and must be
regulated by hepatocyte growth factor activator inhibitors already
during early development. Surprisingly, however, rather than
being a downstream effector of matriptase function, as previously
established for both mouse and human epidermis ([23,24,25,26],
this study), prostasin acts upstream of matriptase during embryogenesis
and is essential for activation of the matriptase zymogen.
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Embryonic Cell Surface Serine Protease Cascade
Figure 7. Neural tube defects and embryonic lethality in HAI-2–deficient mice are not dependent on PAR-2, and combined PAR-1
and matriptase deficiency does not phenocopy combined PAR-1 and PAR-2 deficiency. (A) Distribution of Spint2 genotypes at E9.5 in
PAR-2-expressing (F2rl1 +/+ or F2rl1 +/2 , blue bars) and PAR-2-deficient (F2rl1 2/2 , green bars) offspring from interbred Spint2 +/2 ,F2rl1 +/2 mice. No
Spint2 2/2 embryos were detected irrespective of PAR-2 expression. (B) Frequency of exencephaly observed in HAI-2 and PAR-2-sufficient
(Spint2 + ;F2rl1 + N = 366), PAR-2-deficient (Spint2 + ;F2rl1 2/2 , N = 164), HAI-2-deficient (Spint2 2/2 ,F2rl1 + , N = 18), and PAR-2 and HAI-2 double-
(Spint2 2/2 ;F2rl1 2/2 , N = 12) deficient embryos extracted at E9.5–E11.5. Loss of PAR-2 activity fails to correct neural tube defects in HAI-2-deficient
embryos. (C) Distribution of St14 alleles at E11.5–15.5 in PAR-1-expressing (F2r +/+ or F2r +/2 , blue bars) and PAR-1-deficient (F2r 2/2 , green bars)
embryos from interbred St14 +/2 ;F2r +/2 mice. Loss of PAR-1 activity does not affect embryonic survival of matriptase-deficient mice. (D) Frequency of
exencephaly (Ex), spina bifida (SB), and curly tail (CT) in E9.5–18.5 embryos with different levels of expression of PAR-1 (F2r + or F2r 2/2 ) and matriptase
(St14 + or St14 2/2 ). A total of 326 embryos were analyzed. Loss of matriptase does not significantly increase the incidence of neural tube defects in
PAR-1-deficient embryos. (E) Comparison of the severity of exencephaly in HAI-2-deficient (Spint2 2/2 , N = 29) and PAR-1 and PAR-2 double-deficient
(F2r 2/2 ;F2rl1 2/2 , N = 39) embryos. 95% of affected F2r 2/2 ;F2rl1 2/2 embryos exhibited exencephaly that was confined to hindbrain region of the
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Embryonic Cell Surface Serine Protease Cascade
cranium (HB only, green bars), with the remaining 5% extending to the midbrain region (MB-HB, blue bars). In contrast, only 10% of exencephalies
observed in Spint2 2/2 -deficient mice were confined to the hindbrain, with 59% extended to midbrain, and 31% to forebrain region (FB-HB, red bars).
(F–G9) Ventral (F and G) and dorsal (F9 and G9) view of non-affected control (F and F9) and affected PAR-1 and PAR-2 double-deficient (F2r 2/2 ;F2rl1 2/
2 ) (G and G9) embryos at E9.5. The initial stages of neural tube closure all appear to be unaffected by the combined absence of PAR-1 and PAR-2. (H–
J) Appearance of control (H) and PAR-1 and PAR-2 double-deficient embryos with exencephaly (I and J) at E14.5. Exencephaly in 95% of the affected
PAR-1 and PAR-2 double-deficient embryos was restricted to hindbrain region (HB, two-sided arrow in I) and extended to midbrain (MB-HB, two-sided
arrow in J) in only 5% of the cases. (K and K9) Ventral (K) and dorsal (K9) view of the macroscopic appearance of HAI-2-deficient (Spint2 2/2 ) embryos at
E9.5. Divergence of neural folds (arrows) and defects in neural tube closure extending from forebrain region to cervix are obvious. Open arrowheads
show normal formation of medial hinge points. (L) Macroscopic appearance of a HAI-2-deficient embryo with exencephaly at E14.5. 90% of embryos
presented with exencephaly that included at least midbrain and hindbrain regions of the developing cranium. (M) Histological appearance (nuclear
fast red staining) of PAR-1 and PAR-2 double-deficient embryo with exencephaly at E9.5. Defined medial (arrow) and dorsolateral (arrowheads) hinge
points are clearly visible. Scale bar: 150 mm.
doi:10.1371/journal.pgen.1002937.g007
This finding is perplexing, as matriptase is well-established to be
able to auto-activate, as most clearly evidenced by the inability of
recombinant matriptase protein with the catalytic triad serine
mutated to alanine (S805A matriptase) to undergo activation site
cleavage [56]. Furthermore, prostasin shows no catalytic activity
towards peptide sequences derived from the prostasin pro-peptide
[57] and no reports of prostasin auto-activation have appeared to
date. Substantiating this finding, however, we found that prostasin
efficiently activated the matriptase zymogen in a reconstituted cellbased
assay. These findings are aligned with a recent study
showing that PAR-2 activation in some cultured cells, caused by
exposure of cultured cells to exogenously added activated
prostasin, was blunted by a neutralizing antibody directed against
matriptase [45], providing further evidence that complex and
context-specific relationships between the two membrane-anchored
serine proteases may exist in vivo. Another important
finding relating to the developmental prostasin-matriptase cascade
identified in this study emanated from our biochemical analysis of
placental tissues, which revealed that activated forms of both
matriptase and prostasin were present in a complex with HAI-1 in
placental tissues. This indicates that the regulation of the prostasinmatriptase
cascade by HAI-1 (and likely HAI-2) may occur by
controlling both prostasin and matriptase proteolytic activity.
Furthermore, as both HAI-1 and HAI-2 are very promiscuous and
display potent inhibitory activity towards a number of trypsin-like
serine proteases in vitro [58,59,60,61,62,63,64], it is throughout
plausible that they may also regulate the activity of as yet
unidentified proteases that act upstream of, downstream of or
between prostasin and matriptase. Such profound complexities in
zymogen activation relationships between trypsin-like serine
proteases and for the promiscuity of their cognate inhibitors have
Table 2. Comparison of morphologic features of neural tube
defects observed in Spint2 2/2 and F2r 2/2 ; F2rl1 2/2 mice.
Spint2 2/2 F2r 2/2 ; F2rl1 2/2
Process
Formation of medial hinge point 100% (28/28) 100% (38/38)
Formation of dorsolateral hinge points 3% (1/31) 97% (28/29)
Completion of C1 fusion 100% (28/28) 100% (66/66)
Completion of C2 fusion 90% (27/30) 100% (66/66)
Extent of neural tube defect
Hindbrain only 10% (3/29) 95% (37/39)
Hindbrain and midbrain 59% (17/29) 5% (2/39)
Forebrain to cervix 31% (9/29) 0% (0/39)
Craniorachischisis 0% (0/29) 0% (0/39)
doi:10.1371/journal.pgen.1002937.t002
long been recognized in the coagulation, fibrinolytic, complement,
and digestive systems. The current findings, thus, serve to
underscore that our knowledge of the molecular workings of
membrane-anchored serine proteases is still fragmentary, due to
their quite recent emergence as a protease subfamily.
The outcome of our epistasis analysis querying the contribution
of proHGF, PAR-2, and ENaC to the prostasin and matriptasedependent
embryonic demise of HAI-1- and HAI-2-deficient mice
also was unanticipated. Each of the three proteins has been
genetically validated as a substrate for either matriptase or
prostasin in developmental or post-developmental processes, has
established functions in embryonic development, and is developmentally
co-expressed with both proteases. Nevertheless, their
genetic elimination failed to prevent or alleviate any of the
abnormalities caused by the loss of HAI-1 or HAI-2. Importantly,
our analysis does not exclude that cleavage of either of the three
proteins must be suppressed by HAI-1 or HAI-2 at later stages of
development that cannot be analyzed by the current experimental
approach. Also, the possibility that the lethality of HAI-1- or HAI-
2-deficient embryos is caused by the simultaneous cleavage of
more than one of these substrates cannot be formally excluded. It
was particularly surprising that the neural tube defects associated
with HAI-2-deficiency were unrelated to either excessive or
reduced (through desensitization) PAR-2 activity, despite the
unequivocal contribution of PAR-2 signaling to neural tube
closure, and the wealth of strong circumstantial evidence that
prostasin and matriptase contribute to PAR-2 activation in this
process [45,65]. Equally surprising in this regard, the combined
loss of PAR-1 and matriptase failed to cause the neural tube
closure defects observed in PAR-1 and PAR-2 double-deficient
embryos, showing that matriptase is not essential for initiation of
physiological PAR-2 signaling during neural tube formation.
Previous analysis has identified five other membrane-anchored
serine proteases and fourteen secreted trypsin-like serine proteases
that are expressed during neural tube formation, some of which
can activate PAR-2 in cell-based assays [43,45]. It is therefore
possible that the prostasin-matriptase cascade does contribute to
PAR-2 activation during neural tube closure, but sufficient
residual activation of PAR-2 by other developmentally coexpressed
serine proteases takes place in its absence to allow for
completion of this developmental process. Nevertheless, the careful
comparison of the morphology of neural tube defects in PAR-1
and PAR-2, and HAI-2-double deficient embryos performed here
revealed distinct differences in terms of their anatomical location
and the stage of developmental failure. Taken together, these data
suggest that promotion of neural tube closure by HAI-2
suppression of the prostasin-matriptase cascade and promotion
of neural tube closure by PAR-1/PAR-2 signaling may be
temporally and spatially distinct morphogenic processes.
In conclusion, this study identifies a prostasin-matriptase cell
surface protease cascade whose activity must be suppressed by
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Embryonic Cell Surface Serine Protease Cascade
HAI-1 and HAI-2 to enable early embryonic ectoderm formation,
placental morphogenesis, and neural tube closure.
Materials and Methods
Mouse strains
All experiments were performed in an Association for Assessment
and Accreditation of Laboratory Animal Care Internationalaccredited
vivarium following Standard Operating Procedures. The
studies were approved by the NIDCR Institutional Animal Care
and Use Committee. All studies were littermate controlled.
Spint1 2/2 , Spint2 2/2 , St14 2/2 , Hgfr 2/2 , F2r 2/2 , F2rl1 2/2 ,
Scnn1a 2/2 , and Prss8 fr/fr mice have been described
[38,40,43,49,66,67,68]. Prostasin-deficient (Prss8 2/2 ) mice were
generated by standard blastocyst injection of C57BL/6J-derived
embryonic stem cells carrying a gene trap insertion in the Prss8 gene
(clone IST10122F12, Texas A&M Institute for Genomic Research,
College Station, TX).
Extraction of embryonic and perinatal tissues
Breeding females were checked for vaginal plugs in the morning
and the day on which the plug was found was defined as the first
day of pregnancy (E0.5). Pregnant females were euthanized in the
mid-day at designated time points by CO 2 asphyxiation. Embryos
were extracted by Caesarian section and the individual embryos
and placentae were dissected and processed. Visceral yolk sacs of
individual embryos were washed twice in phosphate buffered
saline, subjected to genomic DNA extraction and genotyped by
PCR (see Table S1 for primer sequences). Newborn pups were
euthanized by CO 2 inhalation at 0uC. For histological analysis, the
embryos and newborn pups were fixed for 18–20 hrs in 4%
paraformaldehyde (PFA) in PBS, processed into paraffin, sectioned,
and stained with hematoxylin and eosin (H&E), or used for
immunohistochemistry as described below. For histomorphometric
analysis of placental labyrinth, the midline cross sections of
plancetal tissues were stained with H&E and the thickness of the
labyrinth was determined as the maximum perpendicular distance
of fetal vessel from the chorionic trophoblast layer.
Immunohistochemistry
Antigens from 5 mm paraffin sections were retrieved by
incubation for 10 min at 37uC with 10 mg/ml proteinase K
(Fermentas, Hanover, MD) for HAI-1 staining, or by incubation
for 20 min at 100uC in 0.01 M sodium citrate buffer, pH 6.0, for
all other antigens. The sections were blocked with 2% bovine
serum albumin in PBS, and incubated overnight at 4uC with
rabbit anti-human CD31 (1:100, Santa Cruz Biotechnology, Santa
Cruz, CA), goat anti-mouse HAI-1 (1:200, R&D Systems,
Minneapolis, MN), mouse anti-human prostasin (1:200, BD
Transduction Laboratories, San Jose, CA), sheep anti-human
matriptase (1:200, R&D Systems) or ENaCc subunit (1:100,
Sigma-Aldrich, St. Louis, MO) primary antibodies. Bound
antibodies were visualized using biotin-conjugated anti-mouse, -
rabbit, -sheep or -goat secondary antibodies (all 1:400, Vector
Laboratories, Burlingame, CA) and a Vectastain ABC kit (Vector
Laboratories) using 3,39-diaminobenzidine as the substrate (Sigma-Aldrich).
All microscopic images were acquired on an
Olympus BX40 microscope using an Olympus DP70 digital
camera system (Olympus, Melville, NY).
Protein extraction from mouse tissues
Placentae were extracted from embryos at E10.5 or E11.5. The
embryonic portion of each placenta was manually separated from
maternal decidua using a dissecting microscope. The tissues were
then homogenized in ice-cold 50 mM Tris/HCl, pH 8.0; 1% NP-
40; 500 mM NaCl buffer and incubated on ice for 10 minutes.
The lysates were centrifuged at 20,000 g for 10 min at 4uC to
remove the tissue debris and the supernatant was used for further
analysis as described below.
Detection of active matriptase and prostasin in mouse
embryonic tissues
Lysates from two placentae of the same genotype were
combined and pre-incubated with 100 ul GammaBind G
Sepharose beads (GE Healthcare Bio-Sciences, Uppsala, Sweden)
for 30 minutes at 4uC with gentle agitation. The samples were
spun at 5,000 g for 1 min to remove the beads, and the
supernatant was then incubated with 5 mg goat anti-mouse HAI-
1 antibody (R&D Systems) and 100 ul of GammaBind G
Sepharose beads for 3 hours at 4uC. The samples were spun at
5,000 g for 1 min, the supernatant was removed, and the beads
were washed 3 times with 1 ml ice-cold 50 mM Tris/HCl,
pH 8.0; 1% NP-40; 500 mM NaCl buffer. The beads were then
mixed with 30 ul of 16SDS loading buffer (Invitrogen, Carlsbad,
CA) with 0.25 M b-mercaptoethanol, incubated for 5 min at
99uC, and cooled on ice for 2 minutes. The samples were spun at
5,000 g for 1 min and the released proteins were resolved by SDS-
PAGE (4–12% polyacrylamide gel) and analyzed by western blot
using mouse anti-human prostasin (1:250, BD Transduction Labs)
or sheep anti-human matriptase (1:500, R&D Systems) primary
antibodies and goat anti-mouse (DakoCytomation) or donkey antisheep
(Sigma-Aldrich) secondary antibodies (both 1:1000) conjugated
to alkaline phosphatase, and visualized using nitro-blue
tetrazolium and 5-bromo-4-chloro-39-indolyphosphate.
Amiloride injections
Five ug per g of body weight of amiloride (Sigma-Aldrich) in
10% DMSO in PBS was administered to pregnant females by
intraperitoneal injection every 24 hours starting on E5.5. Embryos
were extracted on E9.5 by Caesarian section and genotyped as
described, and scored for neural tube closure defects.
Generation of soluble recombinant wildtype, catalytically
inactive S238A, and V170D prostasin zymogens
The generation of pIRES2-EGFP-prostasin has been described
[26]. Substitution of the native prostasin activation site (APQAR)
by the enteropeptidase-dependent cleavage site (DDDDK), and
either the S238A or V170D point mutations were introduced
using the QuickChange Kit (Stratagene, La Jolla, CA) and the
following primers, respectively: 59-GCTCCCTGCGGTGTGG-
CCCCCCAAGCACGCATCACAGGTGGCAGC-39, 59-GAC-
GCCTGCCAGGGTGACGCTGGGGGCCCACTCTCCTGC-
39, and 59-GGCCTCCACTGCACTGACACTGGCTGGGGT-
CAT-39. Successful mutagenesis was verified by sequencing of
both strands of the resulting cDNA. Expression plasmids carrying
individual mutations were transiently transfected into HEK-293T
cells using Turbofect (Fermentas). The cells were grown for two
days and soluble recombinant prostasin was prepared by
treatment of cells with phosphatidylinositol-specific phospholipase
C (Sigma-Aldrich) as described previously [26].
Determination of enzymatic activity of recombinant
prostasin variants
Recombinant wildtype, V170D Frizzy or catalytically inactive
S238A prostasin zymogen variants were first incubated with 5.1 U
recombinant bovine enteropeptidase (Novagen, Cambridge, MA)
overnight at 37uC in enterokinase buffer (Novagen). Following
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Embryonic Cell Surface Serine Protease Cascade
enteropeptidase removal using the Enterokinase Removal Kit
(Sigma-Aldrich), the protein concentration was estimated by
western blot of serially diluted proteins using a reference with
known protein concentration. For substrate hydrolysis assays, the
activated prostasin variants (62.5 nM) were incubated with the
fluorogenic substrate pERTKR-AMC (50 mM final concentration)
(R&D systems) at 37uC in 50 mM NaCl, 50 mM Tris-HCl
pH 8.8, 0.01% Tween-20 buffer, and the fluorescence was
measured using a Wallac plate reader (Perkin Elmer, Waltham,
MA). Each measurement was performed in triplicate. For serpin
complex formation, prostasin variants were diluted in 50 mM
Tris-HCl, pH 9.0, 50 mM NaCl, 0.01% Tween 20 to a final
concentration of 150 nM, incubated with 250 ng recombinant
human protease nexin-1 (PN-1) (R&D Systems) for 1 h at 37uC,
and analyzed using 12% reducing SDS-PAGE and western
blotting, using a monoclonal anti-prostasin antibody (BD Transduction
Laboratories).
Matriptase activation and SRE–luciferase assay
HEK 293 cells were plated in 24-well plates and grown in
DMEM supplemented with 10% FBS for 24 h. Cells were cotransfected
with pSRE-firefly luciferase (50 ng), pRL-Renilla
luciferase (20 ng), pcDNA 3.1 Par2 (100 ng) (Missouri S&T
cDNA Resource Center) using Lipofectamine and Plus reagent
(Invitrogen), pcDNA 3.1 expression vectors containing wildtype
human matriptase or catalytically dead matriptase (S805A), full
length human HAI-1 [69] and empty pcDNA 3.1 vector to
equalize the total amount of transfected DNA. After 36 h the cells
were serum starved over night and then stimulated with 100 nM
recombinant human soluble prostasin (R&D) or vehicle for 6 h.
Cell were lysed and luciferase activity was determined using the
dual luciferase assay kit (Promega, Madison, WI) according to the
manufacturer’s instructions. Chemiluminiscence was measured
using Microtiter Plate Luminometer (Dynex Technologies,
Chantilly, VA) and the SRE activation was determined as the
ratio of firefly to Renilla luciferase counts. The assay was
performed two times in duplicates.
Supporting Information
Figure S1 (A) Western blot detection of protein nexin-1 (PN-1).
Wildtype zymogen (lanes 1 and 2), activated wildtype (lanes 3 and
4), V170D (frizzy) zymogen (lanes 5 and 6), activated V170D
(lanes 7 and 8), S238A zymogen (lanes 9 and 10), and activated
S238A (lanes 11 and 12) prostasin variants were incubated with
(lanes 2, 4, 6, 8, 10, and 12) or without (lanes 1, 3, 5, 7, 9, and 11)
250 ng of recombinant human PN-1. Position of PN-1, and
predicted position of prostasin/PN-1 complexes (not detected by
anti-PN-1 antibody presumably due to significant molecular
rearrangement of PN-1 in the complex with the protease) are
indicated. Positions of molecular weight markers (kDa) are shown
on left. (B) Western blot detection of active matriptase in the fetal
part of the E11.5 placentas of one matriptase-deficient
(St14 2/2 ;Prss8 +/+ ) (lane 1), three wildtype (Prss8 +/+ and St14 +/+ )
(lanes 2,3, and 4), and three prostasin-deficient (St14 +/+ ;Prss8 2/2 )
(lanes 5, 6, and 7) embryos after anti-HAI-1 immunoprecipitation.
A 30 kDa band representing the active serine protease domain of
matriptase (Mat SPD) was present in extracts from wildtype, but
not in matriptase- or prostasin-deficient placentas. (C) Distribution
of Spint2 genotypes at E9.5 in offspring from interbred Spint2 +/2
breeding pairs treated with the ENaC inhibitor, amiloride, at
E5.5–8.5. No Spint2 2/2 embryos were observed.
(TIF)
Table S1 Sequences of PCR primers used for mouse genotyping.
(DOCX)
Acknowledgments
We thank Dr. Mary Jo Danton for critically reviewing this manuscript.
Histology was performed by Histoserv, Germantown, Maryland, United
States of America.
Author Contributions
Conceived and designed the experiments: RS THB JSG. Performed the
experiments: RS KUS PK NAS SG EC KKH SF. Analyzed the data: RS
KUS PK NAS EC THB KKH. Contributed reagents/materials/analysis
tools: SG LKV EH. Wrote the paper: RS THB.
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PLOS Genetics | www.plosgenetics.org 17 August 2012 | Volume 8 | Issue 8 | e1002937

Paper III
Novel assay for detection of active matriptase
Sine Godiksen 1,2 , and Lotte K. Vogel 2
More authors to be added
1 Department of Biology, University of Copenhagen, Copenhagen, Denmark
2 Department of Cellular and Molecular Medicine, University of Copenhagen, Copenhagen,
Denmark
Manuscript in progress
65

Novel assay for detection of active matriptase
Sine Godiksen 1,2 and Lotte K. Vogel 2
Other authors to be added
1 Department of Biology, University of Copenhagen, Copenhagen, Denmark
2 Department of Cellular and Molecular Medicine, University of Copenhagen, Copenhagen,
Denmark
Matriptase is a member of the family of type II transmembrane serine proteases that is crucial
for development and maintenance of several epithelial tissues. Matriptase is synthesized as a
zymogen that is converted by activation site cleavage to a disulfide-linked active form. This
occurs by autoactivation, which has led to the hypothesis that matriptase functions at the
pinnacle of several protease induced signal cascades involved in epithelial tissue formation and
maintenance. Matriptase can be detected in either its zymogen form or in a complex with its
cognate inhibitor hepatocyte growth factor activator inhibitor 1 (HAI-1) in cells, whereas the
active form has been difficult to detect. In this study, we have established an assay to detect
active matriptase using a peptide substrate-based chloromethyl ketone (Cmk) inhibitor. Our
assay is based on the assumption that matriptase able to react with a Cmk peptide inhibitor is
equivalent to active matriptase. In order to detect covalently Cmk peptide-bound matriptase,
we designed a Cmk peptide inhibitor with a biotin moiety in the N-terminal of the small peptide
that allows for streptavidin pull-down and subsequent analysis by Western blotting. This study
presents a novel assay for detection active matriptase in living cells. The assay can be easily
applied in other cell systems and other species.
Abbreviations used: Cmk, chloromethyl ketone; HAI-1, hepatocyte growth factor activator
inhibitor-1; RT, room temperature; SPD: serine protease domain
Introduction
Matriptase (also known as MT-SP1, epithin, TADG-15 and SNC19) is a member of the matriptase
subfamily of type II transmembrane serine proteases. This protease is expressed in most epithelial
cells and has pleiotropic roles in mouse epithelial homeostasis [1-5]. In humans, mutations in the
ST14 gene expressing matriptase, is the underlying cause of congenital ichthyosis [6-8]. More
recently, matriptase was identified as an initiator of the runaway kallikrein protease cascade
leading to an autosomal recessive form of ichthyosis referred to as Netherton syndrome [9].

Ablation of the ST14 gene in mice shows that the protease is essential for postnatal survival.
Matriptase knock-out mice display defects in epidermal barrier function, hair follicle
development, pro-filaggrin processing, and in thymic homeostasis and die within 48 hrs of birth
[5;10]. Tissue-specific or postnatal ablation of matriptase in mice cause severe organ dysfunction,
generalized epithelial demise, and demonstrate a principal and global function of matriptase in
promoting the formation of paracellular permeability barriers in simple and stratified epithelia [4].
Important knowledge about matriptase has also been derived from ablation of the physiological
inhibitors of matriptase; hepatocyte growth factor activator inhibitor-1 (HAI-1) and -2 (HAI-2).
Genetic inactivation of either HAI-1 or HAI-2 leads to failure of placental labyrinth formation
[11;12]. This defect can be completely rescued by simultaneously reducing or eliminating
matriptase expression [12;13]. HAI-1 deficiency in both zebrafish and chimeric mice leads to fatal
defects in epidermal integrity [14-17]. Simultaneous ablation of matriptase in both model systems
completely eliminates these defects [14;16]. Additionally, ablation of HAI-2 in mice leads to
defects in neural tube closure that are partially rescued by genetic inactivation of matriptase
[12;18].
Deregulated matriptase can promote carcinogenesis, as a modest overexpression of wild-type
(WT) matriptase in the epidermis of transgenic mice is sufficient to induce spontaneous squamous
cell carcinoma formation and to strongly potentiate chemical-induced skin carcinogenesis. The
oncogenic effect is mediated through activation of the c-Met-Akt-mTor pathway by matriptasedependent
activation of the cMet ligand; pro-HGF/SF [5;19;20]. A simultaneous increase in
expression of HAI-1 completely negates the oncogenic effect of matriptase overexpression [19].
In colitis-associated colorectal cancer, matriptase was recently assigned as a critical tumorsuppressor
gene. Selective genetic inactivation of matriptase in the intestinal epithelium lead to a
compromised intestinal barrier function associated with chronic inflammation and finally
formation of colon adenocarcinomas in the mouse gastrointestinal tract [21].
Matriptase is synthesized as an 855 amino acid single chain zymogen that undergoes two
successive cleavages to become active. First, matriptase is cleaved in the sea urchin sperm
protein, enteropeptidase, and agrin (SEA) domain by non-enzymatic hydrolysis and subsequently
proteolytically processed at its canonical activation site motif after Arg614 in the serine protease
domain. The serine protease domain (SPD) remains attached to the stem domain by a disulfide
bridge. Shortly hereafter matriptase is inhibited by HAI-1 which leaves a short window of activity
for the disulfide-linked active form of matriptase [22-24]. Using surface biotinylation, we have
previously shown that matriptase undergoes activation site cleavage on the plasma membrane,
and shortly after matriptase is endocytosed in complex with HAI-1 [25].
Despite our knowledge about the important roles played by matriptase, both under normal
physiological conditions as well as in a range of pathological conditions, our understanding of
matriptase and the mechanisms regulating its proteolytic activity is poor. Many epithelial cells
express matriptase that can be detected either in the zymogen or the HAI-1-complexed form
[1;2;22;23;26]. However, the active non-inhibited form, which is the presumably biological active
form of matriptase, has been difficult to detect.

In the present study, we have established an assay to detect active matriptase based on the
assumption that active matriptase is equivalent to matriptase capable of binding an inhibitor. The
assay employs a chloromethyl ketone (Cmk) peptide inhibitor which combined with Western
blotting specifically detects active matriptase.
Materials and methods
Chromogenic assay
0.2 μM matriptase SPD was prepared with 20 mM HEPES pH 7.4, 140 mM NaCl supplemented
with 0.1% BSA (Sigma-Aldrich) in wells of a 96 well plate. Stocks of varying concentrations (5 nM
or 50 μM) of biotin-RQRR-Cmk peptide (American Peptide) were incubated in the same buffer at
37°C for up to 3 hours. At specific time points 100 μl of inhibitor solution was added to the well
containing the indicated concentration of the active catalytic domain of matriptase [27]. Following
additional 10 min incubation at 37°C 10 μl 6.3 mM chromogenic substrate H-D-Isoleucil-L-prolyl-Larginine-p-nitroaniline
(cat. no. S2288, Chromogenix) was added and substrate conversion was
followed in a standard plate reader by 30 min continuous measurements of the absorbance at
405 nm at 37°C. The rate of substrate turnover was determined from the color development
resulting from a pseudo first order reaction due to a substrate concentration far greater than the
expected nM range of protease. To test pH effects, HEPES was replaced by 20 mM citric acid
buffer pH 6.0.
Cell culture
Caco-2 cells were grown in minimal essential medium supplemented with 2 mM L-glutamine, 20%
fetal bovine serum (Gibco), 1 x non essential amino acids, 100 units/ml penicillin and 100 μg/ml
streptomycin (Invitrogen) at 37°C in an atmosphere of 5% CO 2 . For all experiments, 1-2 × 10 6 cells
were seeded into 35 mm tissue culture plates or 0.4 μm-pore-size 24 mm Transwell® filters
(Corning) allowing separate access to the apical and the basolateral plasma membrane. The cell
culture medium was changed every day. Cells were grown until day 11 days post confluence, as
indicated by the tightness of the cell monolayer for filter grown cells, before they were used in
experiments. The tightness of filter-grown cells was assayed by filling the inner chamber to the
brim and allowing it to equilibrate overnight.
Isolation and short-term culture of primary keratinocytes from newborn mice
Epidermis was isolated from newborn mice (d1-2) and grown in culture as described in [28]. In
short, newborn pups were euthanized by decapitation and the torso was submerged in betadine
and ethanol to sterilize the skin. The skin was incubated in 0.25% trypsin w/o EDTA (Sigma-

Aldrich) o/N at 4°C. The dermal portion was discarded and epidermis was minced to release
keratinocytes. The epidermis was resuspended in 45 μM Ca 2+ /10%FBS/Keratinocyte-SFM
(Invitrogen) media and was filtered through a 100 μm cell strainer and centrifuged to remove
stratum corneum pieces. The cell pellet was resuspended in low calcium medium (45 μM
Ca 2+ /Keratinocyte-SFM media) and plated on collagen (BD Biosciences) coated culture plates. Cell
were grown in low calcium medium to sub-confluence and cell culture medium was changed
every second day.
Labeling with biotin-Arg-Gln-Arg-Arg-chloromethyl ketone (biotin-RQRR-Cmk) peptide inhibitor
and S-NHS-SS-biotin
Cells were washed twice; filter grown Caco-2 cells with PBS ++ (PBS supplemented with 0.7 mM
CaCl 2 and 0.25 mM MgCl 2 ) and primary murine keratinocytes with PBS. For labeling of active
matriptase, cells were incubated with 50 μM biotin-RQRR-Cmk (American Peptide) in serum-free
MEM eagle with Earle´s supplemented with 0.2% NaHCO 3 , 100 units/ml penicillin and 100 μg/ml
streptomycin (Invitrogen) at 37°C for the times indicated from the basolateral side. For acidinduced
activation of matriptase, cells were labeled in a physiological phosphate buffer (25 mM
Na 2 HPO 4 , 175 mM NaH 2 PO 4 ) pH 6 or pre-treated with physiological phosphate buffer pH 6 before
labeling in serum-free MEM eagle with Earle´s supplemented with 0.2% NaHCO 3 , 100 units/ml
penicillin and 100 μg/ml streptomycin. As a negative control, cells were labeled with 50 μM of a
corresponding peptide without a CMK group; biotin-Arg-Gln-Arg-Arg (biotin-RQRR). For labeling of
surface proteins, cells were biotinylated from the basolateral side with 1 mg/ml EZ-link Sulfo-
NHS-SS-Biotin (Pierce) dissolved in PBS ++ for 30 min at 4°C. After peptide- and/or ordinary biotinlabeling,
the cells were washed four times with ice-cold PBS ++ . In case of biotin-labeling, residual
biotin was quenched with 50 mM glycine/PBS ++ for 5 min at 4°C and the cells were washed twice
with PBS ++ . Cells were lysed in PBS containing 1% Triton X-100, 0.5% deoxycholate and protease
inhibitors (10 mg/l benzamidine, 2 mg/l pepstatin A, 2 mg/l leupeptin, 2 mg/l antipain, and 2 mg/l
chymostatin). Insoluble material was precipitated at 20,000×g for 20 min at 4°C and the
supernatants were transferred to clean eppendorf tubes.
Streptavidin pull down
Cleared lysates were incubated for 2 hrs with end-over-end rotation at 4°C with 50 μl/24 mm
filter pre-washed streptavidin-coated resin (Pierce), prepared as described by manufacturer. The
streptavidin-coated resin was washed four times with 25 mM TRIS-HCl, 500 mM NaCl, 0.5% Triton
X-100, pH 7.8, and three times with 10 mM TRIS-HCl, 150 mM NaCl, pH 7.8, and biotinylated
proteins were eluted from the streptavidin-coated resin by boiling in SDS sample buffer.
SDS-PAGE and Western blot
Proteins were separated on 10% acrylamide gels and transferred to Immobilon-P PVDF
membranes (Millipore). PVDF membranes were blocked with 10% non-fat dry milk in PBS
containing 0.1% Tween-20 (PBST) for 1 hr at RT and were probed with primary antibody diluted in

1% non-fat dry milk in PBST at 4°C o/N. The next day the membranes were washed 3 times with
PBST, followed by detection of bound primary antibody with horseradish peroxidase (HRP)-
conjugated secondary antibody (Pierce) or alkaline phosphatase (AP)-conjugated secondary
antibody (Sigma-Aldrich). After 3 washes with PBST the signal was developed using the ECL
reagent Super Signal West Femto Maximum Sensitivity Substrate (Pierce) for HRP-conjugated
secondary antibodies according to the protocol supplied by the manufacturer and visualized with
a Fuji LAS1000 camera (Fujifilm Sweden AB) or by nitro-blue tetrazolium and 5- bromo-4-chloro-
3´-indolyphosphate (Pierce) AP-conjugated secondary antibody.
Antibodies
The antibodies used for detection of matriptase in Western blotting were monoclonal mouse antihuman
matriptase antibody M32 that reacts with the A chain recognizing total matriptase as a 70
kDa band (both active and zymogen matriptase in boiled samples) and the 120-130 kDa complex
of matriptase with HAI-1 (only in non-boiled samples) (3ug antibody/blot) [29]; monoclonal
mouse anti-human matriptase antibody M69, which recognizes the 120-130kDa complex of
matriptase with HAI-1 [29]; monoclonal mouse anti-human HAI-1 antibody M19, which recognizes
free 55kDa HAI-1 and the 120-130kDa matriptase-HAI-1 complex [29]; polyclonal rabbit antihuman
matriptase raised against the SPD of matriptase recognizing a 70 kDa band (both active
and zymogen matriptase) under non-reducing conditions and the 70 kDa zymogen form and the
30 kDa protease domain of cleaved matriptase under reducing conditions (Cat. no. IM1014,
Calbiochem). For detection of matriptase in murine cells, sheep anti-matriptase was used (AF3946
diluted 1:1000, R&D). Secondary antibodies include goat anti-mouse HRP-conjugated (10 ng/blot)
(Pierce), goat anti-rabbit HRP-congugated (10 ng/blot) (Pierce) and donkey anti-sheep APcongugated
(10 ng/blot) (Sigma-Aldrich).
Results
Biotin-RQRR-Cmk peptide inhibitor designed to react with active matriptase
In order to detect active matriptase, we designed a peptide inhibitor biotin-RQRR-Cmk consisting
of a tetra peptide; RQRR with an N-terminal biotin moiety and a C-terminal Cmk group. This
inhibitor was designed based on a predicted substrate recognition sequence of matriptase [30].
The Cmk group ensures that the protease-peptide interaction results in the formation of a
covalent bond between the peptide and the protease by alkylation of the active site histidine
attaching a biotin moiety to the active enzyme [31]. The biotin group allows for efficient
concentration of active protease from a complex media by streptavidin precipitation [32].

Biotin-RQRR-Cmk inhibits the proteolytic activity of matriptase SPD in vitro
To verify that biotin-RQRR-Cmk binds matriptase, we tested whether different concentrations of
the peptide inhibitor were able to inhibit the proteolysis of a chromogenic substrate by purified
recombinant matriptase serine protease domain (SPD). 50 μM biotin-RQRR-Cmk renders
matriptase SPD unable to cleave the chromogenic substrate (Fig. 1A, black triangle). Although
chloromethyl ketones are known to be hydrolysed in aqueous solutions with half lives in the order
of 5-20 min [31], we found that an initial concentration of 50 μM biotin-RQRR-Cmk was still
efficient after 180 min pre-incubation at 37°C and able to quench the activity of a sample
containing 0.2 nM matriptase SPD (Fig. 1A, closed circle). A biotin-RQRR-Cmk concentration as low
as 5nM is capable of inhibiting the peptidolytic activity of 0.2 nM matriptase SPD (fig. 1B, black
triangle) even after 60 min of pre-incubation at 37°C (fig. 1B, open circle ), showing that biotin-
RQRR-Cmk is very efficient near-stoichiometric inhibitor of matriptase SPD. Thus, biotin-RQRR-
Cmk is a very efficient inhibitor of matriptase activity and relative stable in aqueous solutions.
Biotin-RRQR-Cmk reacts with a subset of matriptase molecules on the surface of cultured cells
Next, we wanted to test whether the biotin-RQRR-Cmk peptide inhibitor is able to bind active
matriptase on the surface of cells in culture. Differentiated Caco-2 cells have an endogenous
expression of matriptase that can be detected mainly as a zymogen form but also as a two-chain
form in complex with HAI-1 [25;33]. We have previously shown that activation site cleavage of
matriptase takes place on the basolateral plasma membrane of 11 days post-confluent Caco-2
cells [25], indicating that these cells contains active matriptase on the basolateral plasma
membrane.
11 days post-confluent filter grown Caco-2 cells were treated with biotin-RQRR-Cmk from the
basolateral side at 37°C for 2-180 min. Next, cells were lysed and biotin-RQRR-Cmk-labeled
proteases were extracted from cleared lysates using streptavidin-coated resin. Proteins were
released from the resin by boiling in SDS sample buffer. Due to substrate overlap of matriptase
with other trypsin-like serine proteases [30], specificity of the assay is obtained by Western blot
analysis using a matriptase specific antibody. To assess the steady state level of total surfaceassociated
matriptase, parallel cultures were surface-biotinylated with S-NHS-SS-biotin at 4°C.
Matriptase binds biotin-RQRR-Cmk as demonstrated by Western blotting (Fig. 2, lanes 4-6), and
no matriptase could be detected when labeling with a control peptide; biotin-RQRR (Fig. 2, lane 2:
CTRL). Biotin-RQRR-Cmk labeling displayed time dependence at 37°C as an increasing amount of
matriptase was detected with increasing time of incubation with biotin-RQRR-Cmk, indicating that
active matriptase is continuously generated. Comparison of steady state surface-associated
matriptase by standard biotinylation techniques to the accumulated biotin-RQRR-Cmk labeled
matriptase shows that only a fraction of surface-associated matriptase on Caco-2 cells is able to

ind biotin-RQRR-Cmk (compare lane 2 to lanes 3-6). Incubation with biotin-RQRR-Cmk for 180
min at 4°C gave very low signals (data not shown) indicating that the steady state level of
matriptase that is able to bind to biotin-RQRR-Cmk is low. Biotin-RQRR-Cmk also react with other
serine proteases, as prostasin could be detected in Western blotting (data not shown),
emphasizing that specificity of the assay depends on the antibody used for the Western blot
analysis.
Biotin-RRQR-Cmk does not react with the matriptase-HAI-1 complex
After activation site cleavage, matriptase forms a reversible, but SDS-resistant complex with HAI-1
that can be analyzed by SDS-PAGE under non-reducing conditions. In order to investigate whether
biotin-RRQR-Cmk binds to the matriptase-HAI-1 complex, we took advantage of the fact that cells
exposed to slightly acidic conditions has been reported to rapidly convert matriptase from the
zymogen form into the matriptase-HAI-1 complex [24;26;34]. First, we tested whether exposure
to slightly acidic conditions also in Caco-2 cells converts zymogen matriptase into a complex of
activation site cleaved matriptase inhibited by HAI-1.
11 days post-confluent filter grown Caco-2 cells were treated with physiological phosphate buffer
pH 6.0 for 20 min (Fig. 3A, lanes 2, 4, 6, and 8) or left untreated (Fig. 3A, lanes 1, 3, 5, and 7). The
two different lysates were investigated by Western blotting under non-boiled and non-reduced
conditions (Fig. 3A, lanes 1-4 and 7-8) and under reducing conditions (Fig. 3A, lanes 5 and 6) using
different antibodies. Untreated Caco-2 cells mainly contain 70 kDa zymogen matriptase (Fig. 3A,
lanes 1 and 5) and to a minor degree the 130 kDa matriptase-HAI-1 complex (Fig. 3A, lane 1),
whereas matriptase is detected as the 130 kDa matriptase-HAI-1 complex and to a minor degree
70 kDa zymogen matriptase under non-boiling conditions after treatment with physiological
phosphate buffer pH 6.0 (Fig. 3A, lane 2). The generation of the pH 6.0 induced 130 kDa
matriptase-HAI-1 complex is confirmed by antibodies against the matriptase-HAI-1 complex (M69)
(Fig. 3A, compare lanes 3 and 4) and a HAI-1 antibody that also recognizes the 130 kDa
matriptase-HAI-1 complex (Fig. 3A, compare lanes 7 and 8). Analysis of the same samples under
reducing conditions confirms the pH 6.0 induced activation site cleavage of matriptase, as
matriptase was primarily detected as a band migrating at 30 kDa representing the released
disulfide-linked SPD of matriptase after this treatment (Fig. 3A, lane 6). Thus, treatment with
physiological phosphate buffer pH 6.0 induces activation site cleavage of matriptase and
subsequent matriptase-HAI-1 complex formation in Caco-2 cells.
Active matriptase has been shown to have pH optimum at pH 9 [30], we therefore tested the
activity of recombinant matriptase SPD and the inhibitory capacity of biotin-RQRR-Cmk at pH 6 in
the chromogenic assay previously described in Fig. 1. Matriptase SPD is able to cleave the
chromogenic substrate at pH 6.0, although at a much lowered level, as compared to neutral pH
(compare Fig. 3B, square to Fig. 1, square). When 50 μM biotin-RQRR-Cmk was added to the
reaction, no cleavage of the substrate was observed (Fig. 3B, black triangle). Thus, matriptase SPD

is able to bind biotin-RQRR-Cmk at this lowered pH. This enabled us to test whether biotin-RRQR-
Cmk binds to the matriptase-HAI-1 complex.
To examine whether biotin-RQRR-Cmk reacts with the matriptase-HAI-1 complex, we employed
physiological phosphate buffer pH 6.0 to induce matriptase activation. 11 days post-confluent
filter grown Caco-2 cells were treated with 50 μM biotin-RQRR-Cmk from the basolateral side at
37°C for 30 min under different conditions; labeling with biotin-RQRR-Cmk at pH 7.4 (Fig. 4, lanes
3 and 7), labeling with biotin-RQRR-Cmk in physiological phosphate buffer pH 6.0 (Fig. 4, lanes 4
and 8), or pre-treatment with physiological phosphate buffer pH 6.0 for 30 min, before labeling
with biotin-RQRR-Cmk at pH 7.4 for 30 min (Fig. 4, lanes 5 and 9). Additional controls were lysates
of untreated cells (Fig. 4, lane 1) and cells treated with physiological phosphate buffer pH 6.0 (Fig.
4, lane 2) that show pH 6.0 induced matriptase activation and complex formation with HAI-1 (Fig.
4, compare lane 1 and 2). In all cases, aliquots of the total lysates (Fig. 4, lanes 1-5) as well as the
boiled streptavidin pull downs were analyzed in Western blot analysis (Fig. 4, lanes 6-9).
Under the conditions used here, biotin-RQRR-Cmk is unable to dissociate the matriptase-HAIcomplex
formed by pH 6 treatment (Fig. 4, compare lane 9 with lanes 4, 6 and 8). On the other
hand, simultaneous treatment of Caco-2 cells with biotin-RQRR-Cmk and physiological phosphate
buffer pH 6.0 hinders complex formation between matriptase and HAI-1 (Fig. 4, lanes 4 and 5) and
instead more matriptase binds biotin-RQRR-Cmk as detected by Western blot analysis (Fig. 4,
lanes 7 and 8). This observation is supporting the mutual sterical block of the active site in a
competitive manner by either covalent binding to biotin-RQRR-Cmk or non-covalent, SDSresistant
complex formation with HAI-1 [31;35]. Thus, HAI-1 and biotin-RQRR-Cmk competes for
binding to active matriptase.
Biotin-RRQR-Cmk reacts with zymogen matriptase
CMK peptides have been widely used in studies of serine proteases and in most cases Cmkpeptides
react only with the active form of the protease and not with the zymogen form except in
a few cases of autoactivating proteases that have an intrinsic activity enabling the zymogen to
reacts with the Cmk-peptide, one example is tPA [36;37]. Matriptase is also an autoactivating
serine protease implying that the zymogen form has an intrinsic activity [26;38]. To investigate for
an intrinsic activity of matriptase, we labeled 11 days post-confluent filter grown Caco-2 cells with
biotin-RQRR-Cmk from the basolateral side for 180 min at 37°C. Biotin-RQRR-Cmk labeled
proteases were precipitated with streptavidin-coated resin, and subjected to reducing SDS-PAGE.
Samples were analyzed by Western blot analysis using an antibody against matriptase SPD. Under
reducing conditions, matriptase could be detected both as the 70 kDa form (representing
zymogen matriptase) and the 30 kDa form (representing activation site cleaved matriptase) with a
stronger signal intensity of the 70 kDa form (Fig. 5, lane 2). No matriptase could be detected when
labeling Caco-2 cells with a control peptide; biotin-RQRR (Fig. 5, lane 1: CTRL). Thus, both

activation site cleaved and zymogen matriptase is able to bind biotin-RQRR-Cmk indicating that
zymogen matriptase has an intrinsic activity.
Biotin-RRQR-Cmk also reacts with matriptase in other cell systems
The described assay may easily be modified to detect active matriptase from other species as the
specificity of the assay depends on the antibody used. To verify this, the presence of active
matriptase was investigated in cultured primary murine keratinocytes. Keratinocytes from the
skin of matriptase wild-type (WT) and matriptase deficient (KO) newborn pups were isolated,
plated on collagen coated plastic and labeled with 50 μM biotin-RQRR-Cmk at subconfluence for
180 min at 37°C or labeled with S-NHS-SS-biotin to assess surface-associated matriptase. As a
negative control, keratinocytes were labeled with biotin-RQRR. Equal aliquots of cleared lysates
were analyzed by Western blot for total matriptase, prior to streptavidin pull down of biotinylated
proteins. Proteins were released from the streptavidin-coated resin by boiling in SDS sample
buffer and analyzed by Western blotting. By labeling of keratinocytes isolated from mice
expressing WT matriptase with biotin-RQRR-Cmk, we were able to detect active matriptase (Fig.
6, lane 2). No matriptase was detected when labeling with a corresponding control peptide;
biotin-RQRR (Fig. 6, lane 3). No matriptase could be detected in lysates or pull downs of
keratinocytes from matriptase-deficient mice (Fig. 6, lanes 4-6 and 10-12), whereas matriptase
was easily detected in all lysates of the 3 differently treated keratinocyte cultures from mice
expressing WT matriptase (Fig. 6, lanes 7-9). Evaluation of total surface biotinylation and biotin-
RQRR-Cmk labeling of WT murine keratinocytes showed that only a fraction of surface-associated
matriptase on WT keratinocytes cells could be detected by means of biotin-RQRR-Cmk (compare
Fig. 6 lanes 1 and 2). Thus, we have established an assay for detection of active matriptase that
can be easily applied in other cell systems and other species.
Discussion
Matriptase is essential for postnatal survival and tight regulation of matriptase is crucial for a
number of physiological functions including development and maintenance of epithelia [3-5;12-
14;16;17]. Deregulated matriptase activity can also promote skin carcinogenesis [19]. It is
therefore desirable to obtain methods to detect active matriptase to be able to assess active
matriptase under these processes. This has proven difficult as no specific matriptase inhibitor
and/or substrate is commercially available.
For these reasons, we combined antibody specificity with the high affinity of biotin-streptavidin
interaction in the design of a peptide inhibitor-based assay for detection of active matriptase. We

engineered a chloromethyl ketone based tetra-peptide inhibitor that allows for extraction of
inhibitor bound matriptase from complex media by means of an N-terminal biotin moiety.
Specificity of the assay is obtained by Western blot analysis employing specific antibodies against
matriptase. Chloromethyl ketone based inhibitors have been extensively used as inhibitors of
proteases and are active site-directed inhibitors that irreversible bind serine proteases by
alkylation the active site histidine residue of serine proteases rendering it catalytic inactive [31].
The assay is based on the assumption that active matriptase is equivalent to inhibitor bound
matriptase. We show here that matriptase SPD activity is efficiently inhibited by biotin-RQRR-Cmk
and that biotin-RQRR-Cmk is able to label active matriptase on the cell surface of cultured cells.
Upon activation, matriptase rapidly forms a complex with HAI-1 [26;34;39]. HAI-1 is a Kunitz-type
transmembrane serine protease inhibitor that binds to active matriptase in a reversible and
competitive manner as is typical for this family of protease inhibitors [24;40]. The matriptase-HAI-
1 complex comprises a docking of the Kunitz domain I of HAI-1 onto the active site and close
surroundings thereby “capping” the substrate accessibility to the active site of the protease [35].
We found that HAI-1 and biotin-RQRR-Cmk compete for active matriptase and although HAI-1
binds matriptase in an irreversible, but yet very stable manner, biotin-RQRR-Cmk is not able to
dissociate the matriptase-HAI-1 complex under the conditions used in the study. Moreover, the
presence of biotin-RQRR-Cmk hinders matriptase-HAI-1 complex formation.
In most epithelial cell lines and tissues a large fraction of matriptase is found in its zymogen form
[23;26]. In accordance with this, we show that only a fraction of surface-associated matriptase
could be labeled with biotin-RQRR-biotin from cell cultures of primary murine keratinocytes and
unstimulated Caco-2 cells. The zymogen form of trypsin-like proteases is considered to exist in
equilibrium between two conformations, an inactive conformation and an active conformation.
Although most serine proteases are strongly in favor of the inactive form, intrinsic activity has
been reported for a number of autoactivating proteases [36]. Matriptase is an autoactivating
protease implying that the zymogen form has an inherent protease activity. We show that
zymogen matriptase is able to bind biotin-RQRR-Cmk. This finding is supported by a recent study
where a recombinant form of zymogen matriptase in an in vitro assay reacts with an inhibitor
similar to the one used in the present study [41].
Acid-induced activation of matriptase is ubiquitous among epithelial and carcinoma cells and is
followed by rapid complex formation with HAI-1 [24;26;34]. Concurrent with this, we find that
treatment of cells with physiological phosphate buffer pH 6.0 induces matriptase-HAI-1 complex
formation in Caco-2 cells. We also find that a lowering of pH induces a rise in the level of
matriptase that can be labeled with the inhibitor suggesting that there is a brief window between
activation of matriptase and complex formation with HAI-1. This may be of physiological
relevance, as the pericellular environment turns acidic in the transitional layer of the epidermis,
the precise location where matriptase initiates a proteolytic cascade crucial for terminal
epidermal differentiation [3;5;42].
In summary, we have established an assay for detection active matriptase in cell culture. The
availability of an assay for detection of active matriptase can greatly contribute to the further
understanding of the complex biology of matriptase.

Figures
Fig.1 Inhibition of matriptase SPD-mediated chromogenic substrate cleavage by biotin-RQRR-
Cmk. (A) 0.2 μM matriptase SPD was incubated for 10 min at 37°C with (black triangle) or without
(square) 50 μM biotin-RQRR-Cmk before addition the chromogenic substrate to a final
concentration of 300μM. The reactivity of biotin-RQRR-Cmk was tested after 180 min of preincubation
at 37°C (closed circle). Matriptase SPD was unable to cleave the chromogenic
substrate, when biotin-RQRR-Cmk was added (black triangle) even after 180 min pre-incubation
before addition to reaction buffer (closed circle). (B) 0.2nM matriptase SPD was incubated for 10
min at 37°C with (black triangle) or without (square) 5 nM biotin-RQRR-Cmk before addition the
chromogenic substrate to a final concentration of 300μM. The reactivity of biotin-RQRR-Cmk was
tested with 60 min, 120 min, and 180 min of pre-incubation at 37°C before addition to reaction
buffer. 5nM Biotin-RQRR-Cmk is able to completely inhibit substrate cleavage after 60 min preincubation
at 37°C (open circle) and is still able to partly inhibit substrate cleavage after 180 min
pre-incubation at 37°C (closed circle). All measurements were performed in 20 mM TRIS pH 7.4,
140 mM NaCl supplemented with 0.1% BSA at 37°C. The reaction buffer was used as an internal
control to verify that the peptidolytic activity originated from matriptase SPD (grey triangle) and
not unspecific degradation of the substrate over time. Each plot shows the change in optical
density at 405 nm of the reaction mixture as a function of reaction time. The presence of active
protease results in a continued release of a yellow cleavage product resulting in a linear color
development in agreement with a pseudo 1 st order reaction due to the high molar excess of
substrate to protease. Results shown are representative of 3 independent experiments.

Fig. 2. Biotin-RRQR-Cmk reacts with a subset of matriptase molecules on the surface of Caco-2
cells. 11 day post-confluent Caco-2 cells grown on Transwell filters were labeled with 50μM biotin-
RQRR-Cmk from the basolateral side for the times indicated (2-180 min) at 37°C. As a measure of
steady state levels of matriptase, cells were incubated on the basolateral membrane with S-NHS-
SS-biotin at 4°C (lane 1) to biotinylate cell surface proteins. As a negative control, cells were
labeled from the basolateral side with 50 μM control peptide; biotin-RQRR (lane 2). Biotinylated
proteins were precipitated using streptavidin-coated resin and the streptavidin pull downs were
analyzed by non-reducing SDS-PAGE and Western blotting using the monoclonal matriptase
antibody; M32. A tenth of the surface biotinylated sample was loaded (lane 1); whereas total
sample volume was loaded for the other samples (lanes 2-6). Surface biotinylation with S-NHS-SSbiotin
resulted in a band of high intensity (lane 1), indicating that matriptase is abundantly present
on the basolateral membrane. There is a clear time dependence of biotin-RQRR-Cmk binding to
matriptase with weak detection of matriptase after 30 min to the most prominent band after 180
min (lanes 3-6). No matriptase could be detected with the control peptide; biotin-RQRR (CTRL, lane
2). Position of the molecular weight markers (kDa) is indicated on the left. Results shown are
representative of 3 independent experiments.

Fig. 3: Matriptase activation, activity and inhibitor complex formation at pH 6.0. (A) 11 days
post-confluence filter grown Caco-2 cells were either treated at pH 6.0 (lanes 2, 4, 6, and 8) or left
untreated (Lanes 1, 3, 5, and 7) and lysates were analyzed by Western blotting with antibody
against total matriptase (M24; lane 1 and 2), matriptase SPD (IM1014; lanes 5 and 6), matrpitase-
HAI-1 complex (M69; lanes 3 and 4) and HAI-1 (lanes 7 and 8). Samples in lane 1-4, 7 and 8 were
not boiled, while samples in lanes 5 and 6 were boiled and reduced to dissociate the S-S bridged
SPD from the stem domain of activated matriptase. Non-boiled samples of untreated cells reveal
that matriptase is primarily present as a prominent band of 70 kDa and as a minor band at 130
kDa representing the matriptase-HAI-1 complex form (lane 1). Treatment at low pH reveals
matriptase primarily in the 130 kDa complex form and a minor fraction in the 70 kDa form (lane
2). The same rise in matriptase-HAI-1 complex can be detected with the M69 antibody (lanes 3 and
4) and the HAI-1 antibody M19 (lanes 7 and 8). Under reducing conditions, matriptase can be
detected mainly as the 70 kDa form, but also as two band of app. 30 kDa in size (lane 5). These 30
kDa forms are more prominent than the 70 kDa form when cells were treated at pH 6.0 (lane 6).
Treatment with phosphate buffer pH 6.0 and DTT is indicated by +/-. Position of the molecular
weight markers (kDa) is indicated on the left. (B) 0.2 μM SPD were incubated for 10 min at 37°C
with (black triangle) or without (square) 50 μM biotin-RQRR-Cmk before addition the chromogenic
substrate to a final concentration of 300μM. All experiments were performed in 20 mM citric acid
buffer pH 6.0, 140 mM NaCl and 0.1% BSA at 37°C. Matriptase SPD is able to cleave the
chromogenic substrate resulting in color development of the sample at pH 6.0, when biotin-RQRR-
Cmk was omitted (square) and an initial concentration of 50μM biotin-RQRR-Cmk is able to
quench matriptase SPD activity towards the chromogenic substrate also at pH 6.0. A control
containing only buffer verified that the peptidolytic activity originated from matriptase SPD (grey
triangle) and not unspecific degradation of the substrate over time. Results shown are
representative of 3 independent experiments.

Fig. 4. HAI-1 and biotin-RQRR-Cmk competes in inhibition of matriptase
11 days post-confluent Caco-2 cells grown on Transwell filters were labeled with 50 μM biotin-
RQRR-Cmk at neutral pH 7.4 (lane 3 and 7), in physiological phosphate buffer pH 6.0 (Lane 4 and
8), or at neutral pH with a 30 min pre-incubation treatment with physiological phosphate buffer
pH 6.0 (Lanes 5 and 9) for 30 min at 37°C. Samples of lysates were analyzed under non-boiled and
non-reducing conditions (lanes 1-5). Labeled proteases were precipitated using streptavidin
coated resin and released from the beads by boiling. As a negative control, lysate of cells treated
with only physiological phosphate buffer pH 6.0 for 30 min was precipitated (CTRL, lanes 6). The
streptavidin pull downs and non-boiled lysates of the same samples were analyzed by Western
blotting using the monoclonal M32 antibody. Zymogen activation and matriptase-HAI-1 complex
formation is induced with pH 6.0 treatment (lanes 1 and 2, respectively). Likewise, the amount of
matriptase able to bind biotin-RQRR-Cmk also increases after pH 6.0 treatment (compare lanes 7
and 8) and the presence of biotin-RQRR-Cmk hinders complex formation (compare lanes 4 and 5).
Matriptase activation induced prior to biotin-RQRR-Cmk treatment result in matriptase-HAI-1
complex formation (lane 5) and only low levels of matriptase binds biotin-RQRR-Cmk under this
condition (lane 9) comparable to the level detected when treating cells with biotin-RQRR-Cmk at
neutral pH (lane 7). Position of the molecular weight markers (kDa) is indicated on the left. Results
shown are representative of 1 experiment.

Fig. 5. Biotin-RQRR-Cmk inhibits both matriptase zymogen and activation site cleaved
matriptase. 11 days post-confluent Caco-2 cells grown on Transwell filters were labeled with 50
μM biotin-RQRR-Cmk from the basolateral side for 180 min at 37°C. As a negative control, cells
were labeled from the basolateral side with 50 μM control peptide; biotin-RQRR (CTRL), under the
same conditions. Labeled proteases were precipitated using streptavidin-coated resin and the
streptavidin pull downs were analyzed by reducing SDS-PAGE and Western blotting using the
IM1014 antibody raised against matriptase SPD. The biotin-RQRR-biotin peptide labels both the 70
kDa zymogen matriptase and activation site cleaved matriptase, which under these conditions can
be detected as a 30 kDa band. Position of the molecular weight markers (kDa) is indicated on the
left and position of matriptase zymogen and SPD is indicated on the right. Results shown are
representative of 3 independent experiments.

Fig. 6: Detection of active matriptase in cultured primary murine keratinocytes. Murine
keratinocytes were isolated from newborn wild-type (WT) or matriptase-deficient pups and
plated on collagen-coated plastic. The cells were grown until sub-confluence and then labeled
with the S-NHS-SS-biotin (B: lane 1, 4, 7, and 10), with 50 μM biotin-RQRR-Cmk (lane 2, 5, 8, and
11), or with 50 μM control peptide; biotin-RQRR (lanes 3, 6, 9, and 12). Labeled proteases were
precipitated using streptavidin-coated resin, and released from the beads by boiling and analyzed
by SDS-PAGE and Western blotting using the matriptase antibody AF3946. Matriptase is detected
in all lysates of WT keratinocytes (lanes 7-9), whereas no matriptase is detected in the lysates or
pull downs from matriptase-deficient keratinocytes (lanes 4-6 and 10-12). The pull down fraction
of S-NHS-SS-biotin labeled cells (lane 1) as well as the loaded aliquots of WT lysates (lanes 1-3 and
7-9) shows that matriptase is abundantly present at the plasma membrane and that the fraction
of peptidolytic active matriptase only constitute a small fraction of total surface-associated
matriptase (lane 2). Results shown are representative of 2 independent experiments.

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Discussion and perspectives
In the following section the data presented in the three manuscripts enclosed in this thesis will be
discussed starting with a short summary of the key findings
In paper I, we delineate the subcellular trafficking of matriptase, prostasin and HAI-1 to address
how matriptase can act as an upstream activator of prostasin in the epidermis, despite their
different subcellular localizations in polarized epithelial cells. We show that matriptase is routed
to the basolateral plasma membrane, where it is cleaved into its active form. We also found that
both prostasin and HAI-1 are transported to the basolateral plasma membrane from where they
are internalized and transcytosed to the apical membrane. No transcytosis of matriptase could be
detected although matriptase is endocytosed in a complex with HAI-1 from the basolateral
plasma membrane. Interestingly, we could detect active prostasin on both the apical and the
basolateral plasma membrane, as well as active matriptase on the basolateral plasma membrane
by their ability to bind inhibitor-coupled beads. This study propose the basolateral plasma
membrane as the site for matriptase-prostasin interaction and hereby suggest how the apically
located substrate, prostasin, can be cleaved and activated by a basolateral localized protease,
matriptase, in terminal epidermal differentiation.
To gain more knowledge of matriptase-mediated proteolysis in normal physiological processes,
we performed genetic epistasis analyses to identify new component(s) of matriptase-dependent
proteolytic pathways critical to embryogenesis (paper II). We found that a hypomorphic mutation
in Prss8, the gene encoding prostasin, in a similar manner to ablation of ST14, encoding
matriptase, restored nearly all developmental defects associated with HAI-1 and HAI-2 deficiency
indicating that matriptase and prostasin function in the same pathway(s) crucial for survival of
newborn mice. Interestingly, we show that activation of matriptase does not occur in prostasin
deficient placental tissues, whereas activated prostasin is readily detectable in matriptase
deficient placental tissues. Thus, contrary to studies of the epidermis, we demonstrate that
prostasin acts upstream of matriptase and is requires for its activation in placental tissues.
In an attempt to obtain tools for detection of active matriptase, we designed a chloromethyl
ketone peptide inhibitor of matriptase and were hereby able to optimize conditions and establish
an assay for detection of active matriptase in live cell cultures. We found that matriptase is active
on the basolateral plasma membrane by its ability to bind the chloromethyl ketone peptide
inhibitor, biotin-RQRR-Cmk, however only a fraction of total surface matriptase could by extracted
by means of the inhibitor. We show that the chloromethyl ketone peptide inhibitor competes
with HAI-1 for binding to matriptase and that the matriptase zymogen has an intrinsic activity that
enables it to bind the chloromethyl ketone peptide inhibitor. Moreover, we are able to modify the
assay to confirm the presence of active matriptase in cultures of primary murine keratinocytes.
Within recent years, the conception of a matriptase-prostasin proteolytic pathway has been
established in epidermal homeostasis. The existence of such a cascade is based on the facts that
active prostasin is absent in matriptase-deficient epidermis, and that matriptase-deficient mice
85

and mice deficient of prostasin in the skin display nearly indistinguishable phenotypes. Also the
two proteases are co-expressed in a number of epithelial tissues besides the epidermis
[4;9;12;15;16;116].
We have previously delineated the intracellular transport of human HAI-1 recombinantly
expressed in a different model of the polarized epithelia, namely the MDCK cell line. This study
show that HAI-1 has a complex subcellular itinerary and that HAI-1 is transcytosed from the
basolateral plasma membrane to the apical membrane (supplementary I; [27]). The fact that
matriptase in complex with HAI-1 has been purified from human milk, and unpublished results
showing that a 80 kDa protein was co-precipitated with HAI-1 lead us to the hypothesis that HAI-
1, in addition to its protease inhibitor function, could play a role in transporting matriptase in a
matriptase–HAI-1 complex from the basolateral plasma membrane to the apical plasma
membrane, where matriptase could activate prostasin (Sine Godiksen, unpublished results;
supplementary I [27], [44]).
In the current studies (paper I and III), we have used the human Caco-2 cell line as a model for the
polarized epithelial cell. Paper I reports a polarized distribution of endogenous expressed
matriptase, prostasin and HAI-1 in Caco-2 cells, which is in accordance to findings in other cell
types and tissues, showing matriptase and HAI-1 on the basolateral plasma membrane and
prostasin on the apical plasma membrane [9;11;21;23-26]. This makes the Caco-2 cell line a
suitable model system for the studies conducted in paper I and paper III.
The results obtained in paper I also confirmed the previous results obtained in MDCK cells on
subcellular trafficking of HAI-1. However, we were unable to detect transcytosis of matriptase
from the basolateral to the apical plasma membrane as hypothesized, although shedding of HAI-
1-complexed matriptase to the apical media has been observed in polarized Caco-2 cells by us and
others (Stine Friis, unpublished result; [78]).
Instead the results from paper I point to a mechanism where prostasin is routed to the basolateral
plasma membrane followed by transcytosis to the apical membrane as an activated protease.
Thus, the complex itinerary of prostasin, position matriptase and prostasin on the same
subcellular domain and explain how matriptase can activate prostasin in terminal epidermal
differentiation. Whether matriptase act as a direct activator of prostasin in Caco-2 cells, or if other
proteases are responsible for the activation is not shown. On the other hand, our proposed
mechanism for the matriptase-prostasin interaction does not specify any order of the interaction
between the two proteases.
With the generation of matriptase-deficient mice it became clear that matriptase plays a role in
global epithelial development and maintenance [4;5]. Matriptase is believed to be at the pinnacle
of protease cascade(s) because of its ability to autoactivate [28;29;91].
However, it is still unclear which pathways and components that are involved in matriptasedependent
proteolysis. Matriptase and prostasin are co-expressed in a variety of epithelia besides
the epidermis, which has led to speculations of a role for the matriptase-prostasin proteolytic
cascade in other epithelia [9].
In genetic analyses to identify new components of matriptase-dependent proteolytic pathway(s)
necessary for survival of newborn mice, we identified prostasin to be critical to all matriptaseinduced
embryonic defects in both HAI-1 and HAI-2 deficient mice (paper II). Paradoxically, our
study reports that in placental tissue prostasin acts upstream of matriptase and is required for the
86

conversion of zymogen matriptase to active matriptase, which is contrary to what is previously
described for epidermis [15-17;21]. To consolidate this in vivo finding, we show that prostasin is
able to efficiently cleave and activate zymogen matriptase in a recombinant cell based assay.
Interestingly, our data is supported by cell culture studies where addition of active prostasin is
sufficient to activate zymogen matriptase expressed recombinantly in KOLF cells as well as in
HaCaT cells that have an endogenous expression of matriptase [57].
Fig. 13. Outline of matriptase-prostasin interaction in the epidermis and prostasin-matriptase
interaction in placental tissue.
In the epidermis, matriptase acts upstream of prostasin and is required for its action. In placental
tissue the relationship between the two proteases is reversed; thus prostasin is the upstream
protease and required for matriptase activation.
Thus, prostasin may act both as a downstream target but also as an upstream activator of
matriptase in a tissue specific manner, see fig. 13, and/or perhaps constitute a positive feedback
loop similar to the amplification protease cascades operating in coagulation [153]. Further studies
in different epithelial tissues are highly desirable to determine the tissue-specific interplay
between the two proteases, and to identify new components and pathways dependent on
matriptase proteolytic activity in global epithelial homeostasis. It would be interesting to assess
the presence of active matriptase and active prostasin in prostasin-deficient and matriptase-
87

deficient intestinal tissue, respectively, as this is the epithelial tissue to which Caco-2 cells has the
highest degree of resemblance. Still regardless of whether matriptase activates prostasin or vice
versa, the fact that both active matriptase and active prostasin locate on the basolateral plasma
membrane of Caco-2 cells as outlined in paper I can still apply to explain how two proteases with
different steady state localization may interact. Thus it is feasible that in placental tissue,
prostasin could, upon activation on the basolateral plasma membrane, activate matriptase before
being transcytosed to the apical plasma membrane.
The finding that prostasin act upstream of matriptase and is required for its activation is puzzling
in regards to the well-documented ability of matriptase to autoactivate and the fact that prostasin
is incapable of autoactivation due to an unfavorable isoleucine in the P1' position of the activation
cleavage motif of prostasin [107]. It also raises the obvious question of the upstream activator of
prostasin in this setting. Another activator of prostasin is the membrane-anchored serine
protease hepsin [65]. However, hepsin deficient mice exhibit normal embryogenesis and adult
mice display no aberrant phenotype suggesting that another protease is involved in prostasin
activation [154;155]. Establishment of biological functions of the rest of the members of the
family of membrane-anchored serine proteases could provide important information for
determining an upstream activator of prostasin during embryogenesis.
Another interesting aspect and additional layer of complexity of matriptase-prostasin interplay
comes from studies on ENaC activation. Both matriptase and prostasin are able to activate ENaC
in model systems [103;120]. Although membrane anchorage of prostasin is needed for activation
of ENaC in xenopus oocytes, the catalytic activity of prostasin has been reported to be
dispensable for the activational cleavage of ENaC [123;124;156]. This could suggest a role for
prostasin as a co-factor important for the proteolytic activity of another protease towards ENaC.
One could speculate that matriptase participate in prostasin-dependent activation of ENaC. In
paper I we were able to detect matriptase on the apical membrane, and we and others have
observed shedding of matriptase into the apical media, as well as matriptase is present in human
milk (unpublished results, Stine Friis; [44;78]). These findings all indicate that matriptase in fact is
present on the apical plasma membrane. Moreover, by means of the assay established in paper
III, we have shown that active matriptase is present on the apical membrane (unpublished results,
Sine Godiksen). This could position matriptase on the apical plasma membrane to activate ENaC in
a prostasin-dependent manner.
Perhaps prostasin could have a general role as a co-factor, as prostasin has been shown to
enhances matriptase cleavage of EGFR in FT293 cells with no direct cleavage of EGFR by active
prostasin [65]. In this regard, it would also be interesting to examine the necessity for prostasin
catalytic activity in the dependence of GPI-anchored prostasin in plasmin-stimulated ENaC
activation [157]. The role of prostasin as a co-factor could be addressed by the generation of
tissue-specific knock in mouse models of catalytic dead prostasin by mutation of the active site
serine. Thus, the association between matriptase and prostasin is complex and a great amount of
work is needed to unravel the interrelationship of the two proteases in different tissues and cells
types.
88

This thesis also aimed at detecting active matriptase and determining in what subcellular
compartment matriptase is proteolytically processed and activated, and thus able to cleave
downstream substrate(s). In most cell lysates and tissue extracts, matriptase is found in either its
zymogen form or in a complex with its inhibitor HAI‐1 [30;98]. In cell cultures of epithelial and
carcinoma origin, matriptase activation can be induced under slight acidic conditions. This acidinduced
activation of matriptase is fast and efficient and is followed by HAI-1 inhibition within few
minutes [30;31;98;99]. This intimate link between activation and inhibition of matriptase leaves a
small window of action for the protease to cleave its substrates and poses a challenge in detecting
matriptase in its free active form.
Different methods have been used in the three enclosed manuscripts to assess activation of
matriptase and active matriptase. In paper II, we assay for the presence of active matriptase and
active prostasin by co-IP with HAI-1. However, this does not give an assessment of active
protease, but merely inform of whether activation site cleavage has occurred, as we precipitate
HAI-1-complexed prostasin and HAI-1-complexed matriptase. In paper I, we use inhibitor-coupled
sepharose to assess the level of active matriptase. The non-inhibitory LDLR domain of HAI-1 is
important for the matriptase-HAI-1 interaction [28]. For this reason, we used non-endogenous
inhibitors of matriptase and prostasin to avoid possible non-inhibitory interactions and thus to
target only free active matriptase and prostasin. A shortcoming of this procedure is that the
interaction between matriptase and the inhibitor-coupled beads occurs in cell lysates, as
matriptase is sensitive to autoactivation in cell free systems [29].
This prompted us to establish an assay for the detection of active matriptase in live cell cultures
(paper III). The difficulties in setting up a specific assay for the detection of matriptase come from
the great substrate overlap within this family and the absence of a specific inhibitor and/or
specific substrate of matriptase [48;50]. We have based our assay on a chloromethyl ketone
peptide inhibitor with a tetra peptide sequence based on a preferred substrate sequence of
matriptase [50]. CMK peptide inhibitors have long been used in protease research and one of the
great advantages is that they are active site inhibitors and bind proteases in a covalent manner by
alkylation of the active site histidine [158;159]. Although some specificity can be obtained by
altering the peptide sequence of this type of inhibitor, it is very difficult to get absolute specificity
towards a single protease [158]. To circumvent the issue of substrate overlap, we employ specific
antibodies and Western blot analysis in the detection of active matriptase.
We show that active matriptase is present on the basolateral plasma membrane but only
constitute a minor fraction of total surface-associated matriptase although we use vast amounts
of chloromethyl ketone peptide inhibitor (paper III). However, the absence of a linker arm in our
design of chloromethyl ketone peptide inhibitor could cause a reduced affinity for streptavidin
and hereby a reduced detection of biotin-RQRR-Cmk labeled matriptase. For the serine protease
thrombin, a similar assay set-up showed that a linker arm of 7-14 atoms produced the most
sensitive detection in Western blotting [160].
Moreover, our results indicate that biotin-RQRR-Cmk competes with HAI-1 for matriptase binding
however; the precipitation step used prevents us from directly assessing this. Instead, applying
monomeric avidin that bind biotin in a reversible manner would allow us to examine this.
Interestingly, we also find that the zymogen matriptase has intrinsic activity. This is not an
uncommon feature of serine proteases and has been shown for i.a. single-chain tPA [161]. The
89

increase in catalytic activity after zymogen activation varies widely among the different members
of the trypsin protease family, however for single-chain and two-chain tPA the catalytic activities
vary by a factor of only 3-9 [161-165]. It would be interestingly to determine the catalytic capacity
of zymogen matriptase; if strong the zymogen form of matriptase could perhaps be the biological
relevant form in light of the intimate link between activation site cleavage of matriptase and
subsequent inhibition by HAI-1.
Whereas expression studies give valuable insights on matriptase localization and matriptase
mRNA/protein levels, these findings does not reflect the biological active state of the protease.
Having an assay that enables us to assess the level of active matriptase would be valuable in
determining the activation degree of matriptase under different physiological and pathological
states. This is particular the case in assessment of matriptase´s contribution in progression of
cancer. However, applying the assay presented in paper III on tissue extracts would require some
adjustments and also present challenges as matriptase is prone to spontaneous autoactivation in
cell free systems [29]. Additionally it would be desirable with a specific molecular probe for active
matriptase that would also allow visualization of matriptase activity in pathological conditions.
There have been other attempts to identify inhibitors of matriptase. These inhibitors include bisbenzamidines,
sun flower seed trypsin inhibitor-1 derived peptides, antibody-derived inhibitors,
and small molecule peptidyl- derivatives inhibitors [149;166-175]. However, not all inhibitors are
specific for matriptase, e.g. the chemical inhibitor (CVS-3983) of matriptase that was reported to
suppress the prostate cancer growth in nude mice has a high selectivity but is not specific for
matriptase [167]. On the other hand, antibodies constructs have been used for detection of
matriptase positive cancer cells in a mouse xenografts model [166]. Recently an assay was
reported that measured the activity of matriptase on epithelial airway cells. This study use a
different approach than ours and assign proteolytic activity to matriptase by the difference in
measured activity with and without addition of a non-commercial matriptase inactivating
antibody [149]. Even so, despite the methods described above there is still a shortage for specific
biochemical assays for detection of active matriptase.
This thesis reports how complex subcellular trafficking of prostasin enables this protease to
interact with matriptase on the basolateral plasma membrane of polarized epithelial cells and
consolidate how HAI-1 can function as an inhibitor of both proteases. We have shown the
subcellular location for matriptase activation and established an assay for detection of active
matriptase on the surface of living cells. Moreover, this thesis describes a prostasin-matriptase
cascade important for morphogenesis in mice. Together the data presented here has improved
our understanding of matriptase´s role in epithelial physiology.
90

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104

Supplementary I
Hepatocyte growth factor activator inhibitor-1 has a complex subcellular itinerary
Sine Godiksen 1 , Joanna Selzer‐Plon 1 , Esben D. K. Pedersen 1 , Kathrine Abell 1 , Hanne Borger
Rasmussen 2 , Roman Szabo 3 , Thomas H. Bugge 3 , and Lotte K. Vogel 1
1 Department of Cellular and Molecular Medicine, University of Copenhagen, Copenhagen,
Denmark.
2 Department of Biomedical Science, University of Copenhagen, Copenhagen, Denmark
3 Proteases and Tissue Remodeling Unit, National Institute of Dental and Craniofacial Research,
National Institutes of Health, Bethesda, USA
Published in Biochemical Journal, July 2008.
105