restoration manual - Caesar Kleberg Wildlife Research Institute ...

RESTORATION MANUAL
for
Native Habitats of South Texas
Edited by
Paula D. Maywald and Diana Doan-Crider
Caesar Kleberg Wildlife Research Institute
Texas A&M University-Kingsville

An Important Message
from South Texas Natives
The purpose of this manual is to encourage landowners and land managers to restore
native plant communities. Native plants, wildlife, and humans all benefit from restoration
and conservation efforts of good land stewards. Restoration of complex prairie and
native rangeland to a state similar to pre-settlement times is challenging because
vegetation communities and the factors that influence them continually change in time
and space. It is rewarding to watch the landscape change and produce benefits after
restoration has been implemented.
There are two barriers to effective native habitat restoration in South Texas. The first
and most critical is the low commercial supply of locally adapted native seed. Private
landowners, and state and federal agencies are eager to begin using locally adapted
native seed for their re-planting efforts, but very little seed is available in large quantities.
The second is that while some of the methods described in this manual are effective in
other regions, they are not proven planting techniques for South Texas.
In South Texas, unpredictable rainfall is an added challenge. While some variability
is easy to work around, long-term droughts seriously impact restoration efforts and kill
newly planted seedlings or transplanted plants. Keep in mind that rainfall varies even
within the region, and restoration techniques that work well along the Gulf Coast may
not produce results further inland.
Realistically, even the best restoration projects will not duplicate the diversity and
plant species composition of the original native plant communities. Most of these
communities required hundreds or thousands of years to reach their present composition
and structure; if damaged, their restoration will also require some time before they
can function independently. For these reasons, it is important to manage and protect
remaining relic sites in South Texas. Meanwhile, we encourage you to experiment with
techniques and approaches presented in this manual, and to join us in our efforts to
restore native habitats.
Paula D. Maywald, Coordinator
South Texas Natives
Caesar Kleberg Wildlife Research Institute
Texas A&M University-Kingsville
700 University Blvd.
MSC 218
Kingsville, TX 78363-8202
361-593-5550
paula.maywald@tamuk.edu
http://ckwri.tamuk.edu

Table of Contents
Acknowledgements
How to Use This Guide
Introduction
Why Restoration? 1
What are Native Species? 2
About Non-native Plants
The Importance of Native Plant Diversity
Chapter 1
South Texas Ecology and Climate
Rainfall
Temperature
Soils
Habitats
Chapter 2
Water
South Texas Moisture.. or Lack Thereof
Irrigation
Planning around Water
Chapter 3
Planning Your Project
Define Your Goal
Chapter 5
Soils
Importance of Soils to the Restoration Process
Physical Properties
Organic Matter in Soils
Soil Microbial Activity
Chemical Properties
Topsoil
Soil Amendments
Chapter 6
Site Preparation
Timing is Everything!
Site Preparation Methods
Brush Control
Weed and Non-Native Grass Removal
Alleviating Soil Compaction
Cover Crops
Seedbed Preparation
Weed and Non-Native Grass Removal
Alleviating Soil Compaction
Cover Crops
Seedbed Preparation
Chapter 7
Acquiring Seed
Purchasing Seed from Dealers
Collecting Wild Native Seed
Seed Maturity
Chapter 4
Methods of Establishing Native Plants
Cultivating Native Seed
Wild Native Seed
Wild Native Hay
Natural Revegetation
Transplanting
Integrating Methods
ii

Chapter 8
Seed Processing and Storage
Seed Processing
Storage
Short-Term Seed Storage
Long-Term Seed Storage
Protection from Insects
Native Hay Storage
Bulk or Bagged Native Seed Storage
Chapter 9
Seed Application
Broadcast Seeding
Drill Seeding
Seed Drill Calibration
Seeding Cultivated Land
Hydroseeding and Mulching
Wild Harvested Native Hay Application
Appendices
Appendix A - Glossary of Plant Basics
Appendix B - Suggested Plant Species
for Use in Restoration
Appendix C - Selected Plant Profiles
Appendix D - Common and Scientific
Names Mentioned in Text
Appendix E - References & Literature Cited
Appendix F - Consultants
IGNORE PAGE
NUMBERS PAGE
FOR NOW
Chapter 10
Propogating and Transplanting
Why Propagate Native Plants?
Germination
Testing for Germination Rates
Seed Scarification
Planting Containers
Potting Mediums for Containerized Plants
Propagation
Seedling Mainenance
Transplanting Native Plants
Maintenance after Transplanting
Chapter 11
Management Practices after
Restoration
Important Factors to Consider before Grazing
or Burning a Newly Restored Site
Grazing
Prescribed Burning
Grazing Following Controlled Burns
iii

Acknowledgements
Restoration Manual Funding
Funding for the production of this manual was provided by Reliant Energy and Mr. Ken Leonard,
L&H Packing, Inc. We are deeply grateful for their steadfast support.
Founding Donors of South Texas Natives
• Robert J. Kleberg Jr. and Helen C. Kleberg Foundation
• Lee and Ramona Bass Foundation
Principal Sustaining Sponsor of South Texas Natives
• Texas Department of Transportation
Sustaining Donors and Sponsor of South Texas Natives
• Joan and Herb Kelleher Charitable
• ExxonMobile Foundation
iv
Scientific Contributors
• Charity Lawson, Caesar Kleberg Wildlife Research Institute, Texas A&M University-Kingsville
• Fred Bryant, Ph.D., Caesar Kleberg Wildlife Research Institute, Texas A&M
University-Kingsville
• Barrie Cogburn, Texas Department of Transportation
• Timothy E. Fulbright, Ph.D., Caesar Kleberg Wildlife Research Institute, Texas A&M
University-Kingsville
• Wayne Hanselka, Ph.D., Texas Cooperative Extension, Department of Rangeland Ecology and
Management, Texas A&M University
• Brian Hays, Texas Cooperative Extension, Wildlife and Fisheries Sciences Department
• Richard Hoverson, Ph.D., Wildlife Habitat Improvement
• David Mahler, Environmental Survey, Inc.

• Dennis Markwardt, Texas Department of Transportation
• Paula D. Maywald, South Texas Natives, Caesar Kleberg Wildlife Research Institute, Texas
A&M University-Kingsville
• Bill Neiman, Native American Seed
• Bill Ocumpaugh, Ph.D., Texas A&M University Agricultural Experiment Station
• Kerry Olenick, USDA-NRCS
• Alfonso Ortega, Ph.D., Texas A&M University-Kingsville
• Allen Rasmussen, Ph.D., Texas A&M University-Kingsville
• John Lloyd Reilley, USDA-NRCS E. Kika de la Garza Plant Materials Center
Scientific Contributors - continued
• Fred Smeins, Ph.D., Department of Rangeland Ecology and Management, Texas A&M
University
• Forrest Smith, South Texas Natives, Caesar Kleberg Wildlife Research Institute, Texas A&M
University-Kingsville
• Matt Wagner, Ph.D., Wildlife Division, Texas Parks and Wildlife Department
• Steve Whisenant, Ph.D., Department of Ecosystem Science and Management,
Texas A&M University
• Lisa Williams, The Nature Conservancy of Texas
• Larry Zibilske, Ph.D., USDA ARS Kika de la Garza Subtropical Agricultural Research Service
Principal Editors
• Paula Maywald, Coordinator, South Texas Natives, Caesar Kleberg Wildlife Research
Institute, Texas A&M University-Kingsville
• Diana Doan-Crider, Ph.D.
Contributing Editors
• Forrest Smith, South Texas Natives, Caesar Kleberg Wildlife Research Institute, Texas A&M
University-Kingsville
• Charity Lawson, Caesar Kleberg Wildlife Research Institute, Texas A&M University-Kingsville
Layout & Graphic Design
• Diana Doan-Crider, PhD
Prinicipal Photographer
• Forrest Smith, South Texas Natives, Caesar Kleberg Wildlife Research Institute, Texas A&M
University-Kingsville
Contributing Photographers
• Diana Doan-Crider, Ph.D.
• Charity Lawson, Caesar Kleberg Wildlife Research Institute, Texas A&M University-Kingsville
• Cody Lawson
• Paula Maywald, South Texas Natives, Caesar Kleberg Wildlife Research Institute, Texas A&M
University-Kingsville
• Texas A&M University-Kingsville stock photos
Cartoon Graphics
• Justin Hall, Dances for Peas, Inc.

We would like to thank the following for allowing us the use of
excerpts from their publications:
• Bryant, F. C. and T. E. Fulbright - Last Great Habitat, Caesar Kleberg Wildlife
Research Institute Special Publication No. 1, Texas A&M University-Kingsville 2002.
• Native Plant Revegetation Guide for Colorado - Caring for the Land Series Volume III,
Colorado Natural Areas Program, Colorado State Parks, and Colorado Department of Natural
Resources, 1998.
• Pratt, M., and G. A. Rasmussen - Drill Calibration, Utah State University Extension, http://
extension.usu.edu/files/natrpubs/range2.pdf.
• Welch, T. (additional updates by A. McGinty, J. F. Cadenhead, C. W. Hanselka, D. N. Ueckert,
and S. G. Whisenant) - Chemical Weed and Brush Control Suggestions for
Rangeland. Texas Agricultural Extension Service, Texas A&M University System.
• Wagner, M., F. Smeins, and B. Hays - Pastures for Upland Birds: Restoration of Native Plants
in Bermudagrass Pastures.
Updates and Input
The emphasis of this manual is to provide basic understanding of processes involved in establishing
native species in South Texas. It does not contain suggestions for every property, for every possible
revegetation scenario, or plant species native to South Texas. Knowledge regarding restoration
will continue to increase over time through implementation of these strategies and continued
experimentation. We hope this guide serves as a good place to begin, and we will periodically
update the manual as new restoration information becomes available. We look forward to hearing
about your restoration experiences.
Updated addendums will be published as information is accumulated and techniques are refined,
and will be made available in a PDF format compatible with this notebook through our website at
http://ckwri.tamuk.edu. South Texas Natives is also interested in the results of your projects or
research, so please send input that you would like to share to the following address:
Paula Maywald
South Texas Natives
Caesar Kleberg Wildlife Research Institute
700 University Blvd., MSC 218
Texas A&M University-Kingsville
Kingsville, TX 78363-8202
361-593-5550
vi

How To Use This Guide
Before beginning any vegetation restoration project, we recommend that you
first become familiar with this guide. Also, it is important that you work closely
with your local extension agent, range specialist, or plant materials center.
This guide provides information on how to select, plant, and maintain South
Texas native plant species for a wide range of landscaping, revegetation,
and reclamation needs. Instructions for the design, planning, execution,
and maintenance of revegetation projects are also presented. In addition to
the information provided in the text, additional resources are provided in the
appendices.
SPECIAL SYMBOLS AND COLOR-CODES USED IN THIS GUIDE:
Blue Text
Cross-referenced topics
Orange Text
Essential References
Literature Cited boxes Blue Boxes
Useful References boxes Blue-gray Boxes
Helpful Tips boxes
Green Boxes
Seed Head Eddy will be used through out this manual to emphasize special tips.
KEY POINTS FOR A SUCCESSFUL RESTORATION PROJECT
• Know the resource managers in your area. We have included a list of contacts for the South
Texas region in Appendix X.
• Become familiar with additional published sources of information for South Texas such as those
listed throughout this guide, especially the USDA Soil Surveys for your county.
• Use website searches as an information tool, but be prepared to accept and adapt
strategies and techniques to the South Texas region.
Save for last
vii

Notes

Whitebrush (Aloysia gratissima)
INTRODUCTION

Eastern gamagrass (Tripsacum dactyloides)
SOUTH TEXAS NATIVES
INTRODUCTION
South Texas Natives is an initiative to develop and
promote native plants for the restoration and reclamation of
habitats on private and public lands. In a collaborative effort with
the USDA-NRCS E. Kika de la Garza Plant Materials Center, our
goal is to provide economically viable sources of native plants and
seeds to both the private and public sector for the restoration of native
plant communities in South Texas. Our objectives include developing
and implementing strategies to establish native seeds and plants while
minimizing the influence of introduced plants upon native habitats.
Habitat deterioration is generally caused by human and/or animal activity, and
can be augmented by external stresses such as droughts or disruption of natural
fire patterns by human activities. Altered water cycles are major ecosystem
dysfunctions on semi-arid South Texas rangelands. Overgrazing during the past
250 years, combined with a reduction in naturally occurring fires and frequent
droughts, disrupted the ecological processes of nutrient cycling, energy
flow, plant community dynamics, and in particular, hydrologic
Why Restoration? processes. Changes in soil fertility, increased erosion, and
compaction contribute to decreased water infiltration and
increased runoff, resulting in lower vegetation diversity, reduced surface coverage, increases in
brush, and lower productivity. Over time, plant species may lose vigor and die. As herbaceous
biomass decreases and bare ground increases, ecological processes of water, nutrient, and energy
cycling are disrupted. This results in lost capacity for the plant community to maintain itself and
further deterioration occurs. After original plant communities have been severely disturbed, stable
processes have been upset, and invader plants have become established, the plant community
cannot easily or economically be restored to simulate its original state.
The Society for Ecological Restoration defines restoration as “the process of assisting the recovery
of an ecosystem that has been degraded, damaged, or destroyed.” Restoration is a holistic
process which not only involves revegetation, but also entails the removal of non-native species,
the reintroduction of soil biota (such as invertebrates, insects, and fungi), and the implementation of
management strategies that will help the system function in a healthy manner. Increasing stability
and water infiltration in the soil surface initiates repair and maintenance of damaged processes
that enhance plant production and protect the soil surface with plant litter or living vegetation.
What are Native Species?
A native species is “a plant species indigenous to the natural plant community where it is found.”
For the purposes of this handbook, the terms alien, non-native, and exotic describe plants that
have been introduced into the 33 county region of South Texas. To keep things simple, this guide

uses “non-native” throughout the text. In addition, the term “ecotype” is used to describe a subset
(or population) of a native plant species adapted to a specific (or localized) set of environmental
conditions.
When settlers first entered Texas in the mid 1700s, they brought seeds of plants from Europe
and the Mediterranean. Some were seeds planted in the Old World for food crops, windbreaks,
landscaping, erosion control, and livestock forage. Many other seeds arrived accidentally, mixed
with crop seeds, hay, animal feed, or even in the ballast of ships. These came westward with
shipments of agricultural goods, or they were dispersed along irrigation ditches, railroad tracks,
wagon roads, and cattle trails. Today, either by accident or design, introduction of non-native
plants into Texas continues. Because of past and present human activities, non-native species
have taken over much of South Texas’ landscape.
About Non-native Plants
Many non-native plants spread rapidly and displace native species. Native plant species have
evolved with local environmental factors, such as insects, diseases, and herbivores that regulate
their population numbers. In most cases, non-native species have not evolved with these same
factors. If left uncontrolled, some non-native species can form extensive monocultures and reduce
the diversity of native plant communities. This can be detrimental because many non-native
species do not provide adequate habitat for native wildlife and insects. It is important to note that
not all non-native plants are invasive, including food crops and landscape plants.
Why are native plants important to South Texas?
Concerns about conservation of tropical rainforests and other well-known regions of the world
are widely publicized, yet a region of inestimable biological wealth lies relatively unrecognized
on the back doorstep of North America. The region lying south of a line from Port O’Connor to
Victoria, northwest to San Antonio and west to Del Rio known as
“South Texas” is one of the most biologically diverse regions
in the world. In fact, it is termed “hyper-diverse” by
many ecologists, and is considered by some as one
of “the last great habitats” remaining intact in North
America.
The diversity of native South Texas habitats ranges
from the fine sands of the Coastal Sand Plain to the
caliche ridges of the Bordas Escarpment; and from the
riparian woodlands of the Nueces River to the shrublands
of the Rio Grande Plains. This diversity supports a wide array
of wildlife species, ranging from migratory birds such as sandhill
cranes and piping plovers, to more permanent residents such as ocelots
and white-tailed deer.
An “invasive
species” is defined as a
species that is 1) non-native (or alien)
to the ecosystem under consideration and
2) whose introduction causes or is likely to
cause economic or environmental harm or harm
to human health (definition from Executive
Order 13112). A plant may be invasive
under certain soil and/or environmental
conditions, but not others.
Native plants are intrinsic to the overall resilience and stability of this unique
region, and are a critical component of the numerous food and energy cycles

that maintain this biological diversity. Establishment and restoration efforts using native South
Texas plants will help maintain the region’s important phytogenetic resources and the ecosystems
that are part of South Texas’ biological heritage.
The Importance of Native Plant Diversity
Plant community diversity is important to wildlife and water resources. Plant species vary in timing
of growth (e.g., cool versus warm season) and timing of seed and fruit production. In habitats with
a large number of different plants, plants or groups of plants produce food for herbivores at differing
times of the year. In habitats with low plant species diversity, odds are greater that food will be
absent at some time of the year. Where plant diversity is high, for example, white-tailed deer are
able to maintain a more constant level of nutrition throughout the year.
Species-rich plant communities are more resistant to drought than species-poor plant communities,
an important attribute in semiarid habitats. Resilience, or rate of return to pre-drought conditions,
is also greater for more species-rich communities. Species-rich communities are commonly more
biologically productive than species-poor communities.
USEFUL REFERENCES
National Invasive Species Information Center - http://www.invasivespeciesinfo.gov
Society for Ecological Restoration International - http://www.ser.org
Texas prickly pear on South Texas Coastal Prairie

Notes

South Texas Sunrise
SOUTH TEXAS ECOLOGY
AND CLIMATE

SOUTH TEXAS ECOLOGY
AND CLIMATE
Rainfall
The extraordinary diversity of plants, wildlife, and habitats is partly
driven by an environment that is quite variable among years
and across the South Texas landscape. Vegetation in South
Texas is exceptionally resilient to heat and low rainfall, and is
able to make a noticeable comeback even after experiencing
years of severe drought. Across the region, the annual
rainfall averages 24.5 inches, but fluctuations in South Texas
precipitation can be dramatic. Between the years 1900 and
1983, the driest year was 1917, when the regional rainfall
average was 9.5 inches. The wettest year was only 2 years
later in 1919, however, when the regional average was 40.8
inches. Overall, 36% were drought years and 34% were wet years 1 .
Temperature
Spanish Dagger
(Yucca treculeana)
South Texas’ climate is subtropical subhumid-to-semiarid, with high temperatures, high evapotranspiration
rates, and very few killing frosts 1 . The average annual air temperature in South Texas
exceeds 70°F, which is comparable to southern Florida. July temperatures commonly exceed
98°F, and these extremely warm temperatures have a profound impact on the ecology of plants and
animals. Woody plants, for example, play a keystone role in the region’s ecology by moderating
the thermal environment beneath their canopies. This, in turn, provides protective cover for many
plant and animal species in the region.
Spring thunderstorm, Jim Hogg Co.,TX

Average Annual Precipitation
Soils
Soils in South Texas cover the entire spectrum of particle sizes, ranging from coarse sands of the
Ingleside Prairie to fine montmorillonitic clays of the mid and lower Coastal Prairie. The Coastal
Sand Plain is characterized by deep, white-colored, aeolian fine sands. Active, blowing dunes
occupy about 5% of the Coastal Sand Plain. These dunes migrate across the landscape in the
direction of the prevailing winds from southeast to northwest and create unique, “mini-desert”
microhabitats that add to the diversity of the surrounding landscape.
Sandy loam soils are common in the western South Texas Plains. Many of the upland sandy loam
soils in western South Texas have a reddish hue. The “red sands” are renowned for supporting
productive wildlife habitat.
Saline soils of the Maverick, Montell, and Monteola soil series cover thousands of acres in the
western part of South Texas 2 . These soils support plants such as saladillo and armed saltbush,
which are known as “halophytes.” Halophytes are plants possessing unique characteristics that
enable them to survive in the harsh environment of saline soils. For example, some halophytes
exude excess salt on the surface of their leaves to prevent too much salt from accumulating inside
the leaves. Leaves of succulents such as Texas varilla swell because they absorb water to prevent
salt from becoming too concentrated in their tissues.

Soils from a Ranch in Dimmit County, Texas
The Bordas escarpment consists of shallow soils underlain by a thick layer of caliche. Oriented
along a generally north to south axis, the rolling hills of the escarpment bisect South Texas. The
thin soils of the escarpment are high in calcium and support a diverse and unique assemblage
of woody plant species. These soils are not amenable to cultivation, but support native plants
that provide excellent wildlife habitat. A great variety of soil types can exist in just one locale, as
illustrated by a ranch in Dimmit County (figure above). The variety of soils and the resulting variety
of habitat types found in relatively small geographic areas within South Texas are major factors
affecting the high species diversity of plants and animals in this region of Texas.
Driven by variability in soils, precipitation, and temperature, South Texas has a mixture of subtropical,
eastern deciduous forest, and Chihuahuan desert plant and animal species. There are 1,411
species and subspecies of vascular plants in the Texas Coastal Bend region, within a mere 50-
65 mile radius of Corpus Christi, Texas. Different combinations of plant species create an array
of communities ranging from wetlands to sand dunes, and from grassland to oak woodland and
semiarid shrubland.
Habitats
Freshwater Wetlands
Spiny aster and longtom paspalum are common
dominant plant species of periodically inundated
areas in eastern South Texas 4 . Coontail, water
nymph, water stargrass, wigeongrass, sago
pondweed, and muskgrass are common aquatic
plants in submerged freshwater communities of
ponds and lakes. Floating-leaved plants include
lotus and other species within the same family.
Marsh edges are dominated by bulrushes, cattails,
and sedges. Edges of lakes and ponds support
stands of longtom and clubhead cutgrass.
Northern shoveler (Anas clypeata) in a freshwater
wetland, Nueces Co., TX

Laguna Madre
The Laguna Madre of South Texas and Laguna
Madre del San Antonio in northern Mexico together
form one of only 3 to 5 hypersaline lagoonal areas
in the world of significant size 3 . The Laguna
Madre supports many of the seagrasses along
the Texas Coast; these plants play a critical role in
the reproductive cycles of many estuarine fish and
invertebrates by providing refuge or habitat during
at least part of their life cycle. Recent increases
in disruptive human activity have resulted in
reductions in water clarity which, in turn, may be
Laguna Madre, Kleberg Co., TX
affecting seagrass cover. In addition, recent dredging activity has altered the salinity of the lower
Laguna Madre, which has caused a shift in seagrass composition.
Shrublands
There are 6 shrubland associations in South Texas 6 . The honey mesquite-granjeno association is
the major shrubland association in eastern South Texas. Poorly drained soils support an association
dominated by huisache and prickly pear. Saline or sodic soils are characterized by an association
of stunted honey mesquites and pricklypear. The caliche hills of the Bordas Escarpment, which
extends from Starr County north and eastward to the Nueces River, are dominated on top by
the guajillo-ceniza association and on the upper slopes by the blackbrush acacia-twisted acacia
association. The creosotebush-pricklypear association is found on gravelly soils in western South
Texas, and is the driest vegetation association in the region. It contains numerous plant and animal
species characteristic of the Chihuahuan Desert of western Texas and northern Mexico.
Prairies
Prairies are a vanishing vegetation type in Texas. Much of the original Shortgrass Prairie in the Texas
Panhandle has been converted to farmland. Natural vegetation of the Blackland Prairie, Coastal
Prairie, and Fayette Prairie has been almost completely lost to cultivation and development 5 . Less
than 5% remains of the original Tallgrass Prairie that once extended from Oklahoma south through
the eastern half of Texas to the Gulf Coast.
10
Honey mesquite (Prosopis glaudulosa) - Granejo (Celtis pallida) association, Atascosa Co., TX

South Texas is rich in native prairies with at least 8 distinct native prairie types, including the
Fayette Prairie, upper Coastal Prairie, lower Coastal Prairie, Ingleside Prairie, Kenedy Sand Prairie,
Bluestem-Sacahuista Prairie, Sea Oats Prairie, and Southern Cordgrass Prairie 6,7,8 . The Fayette
Prairie is similar to the Blackland Prairie, which is dominated by little bluestem 7 . The Fayette Prairie
originally paralleled the Coastal Prairie extending southwest almost to the Frio River. The lower
Coastal Prairie is dominated by little bluestem and multiflowered false rhodesgrass and extends
from Kleberg County northward to the San Antonio River where it merges with the upper Coastal
Prairie 6,9 . The Ingleside Prairie lies along the Laguna Madre from Refugio County to the mouth
of Baffin Bay. Because of land ownership patterns, the portion of the Ingleside Prairie between
Corpus Christi and the mouth of Baffin Bay remains largely intact. Dominant grasses of Ingleside
Prairie are seacoast bluestem, switchgrass, and fringeleaf paspalum 7 .
The Kenedy Sand Prairie, found in the region extending from the mouth of Baffin Bay south to
Willacy County and west to Jim Hogg County, is the largest remaining, intact prairie in Texas.
Seacoast bluestem is the prevailing dominant plant species, with gulfdune paspalum dominating in
swales and moderately drained flats in the eastern part of the region 10 .
The Bluestem-Sacahuista Prairie occurs in a belt from 50 to 150 miles inland along the Gulf Coastal
Plain 8 . Dominant species include little bluestem, seacoast bluestem, and gulf cordgrass. The
barrier islands off the southern coast of Texas support the Sea Oats Prairie dominated by seacoast
bluestem, seaoats, and gulfdune paspalum. The Southern Cordgrass Prairie is found in a narrow
band adjacent to the Gulf Coast in both freshwater and brackish marshes. Dominant species
include smooth cordgrass and marshhay cordgrass.
Mesquite-bluestem prairie, Jim Hogg Co., TX
11

Woodlands-Forests Along Rivers and Streams
Woodlands found in riparian habitats in South Texas are dominated by sugar hackberry and
huisache 6 . Other trees, which rise to dominant status depending on location, include eastern
cottonwood, post oak, live oak, cedar elm, anaqua, honey mesquite, pecan, black hickory, shagbark
hickory, Texas persimmon, Texas ebony, mustang grape, and muscadine grape.
Uplands are often veined with thin riparian areas known as ramaderos 11 . Ramaderos receive runoff
water from adjacent uplands and support comparatively lush vegetation. These areas are critical
nesting, feeding, and loafing areas for wildlife. Ramaderos also serve as corridors for animal
movements providing a linking network with surrounding habitats. Thus, they are critical areas for
maintaining biodiversity in the surrounding landscape. Sadly, more than 90% of the riparian habitat
in South Texas has been cleared for agricultural or urban use. Conservation of remaining riparian
areas is critical to prevent further loss of biodiversity.
Sandy soils in South Texas support live oak-post oak woodlands 6 . A post oaklive
oak/little bluestem community is the major woodland community in a belt
south of San Antonio in the northeastern portion of South Texas. A live oakhoney
mesquite/seacoast bluestem community occupies parts of the Coastal
Sand Plain interspersed within the Kenedy Sand Prairie.
Live Oak Woodland, Kenedy Co., TX
12
Engelmann’s daisy (Engelmannia pinnatifida)

Last Great Habitat Plant Communities
Map of broad-scale plant
communities in South Texas.
Modified from T. McLendon,
1977, Texas A&I University,
Kingsville, Texas, unpublished.
LITERATURE CITED
1
Norwine, J., and R. Bingham. 1986. Frequency and severity of droughts in South Texas: 1900-1983. Pages 1–17 in
Livestock and wildlife management during drought (R. D. Brown, editor). Caesar Kleberg Wildlife Research
Institute, Kingsville, Texas.
2
Fanning, C. D., C. M. Thompson, and D. Isaacs. 1965. Properties of saline range soils of the Rio Grande Plain.
Journal of Range Management 18:190-193.
3
Gunter, G. 1967. Vertebrates in hyper saline water. Contributions in Marine Science 12:230–241.
4
Drawe, D. L., A. D. Chamrad, and T. W. Box. 1978. Plant communities of the Welder Wildlife Refuge. Welder Wildlife
Foundation, Contribution Number 5, Series B, Revised. 38pp.
5
Diamond, D. D. 1990. Almost gone…almost forgotten. Texas Parks and Wildlife Magazine 48(6):13–16.
6
McClendon,
T. 1991. Preliminary description of the vegetation of South Texas exclusive of coastal saline zones. Texas
Journal of Science 43:13–32.
7
Johnson,
M. C. 1963. Past and present grasslands of southern Texas and northeastern Mexico. Ecology 44:456–
466.
8
Shiflet,
T. N. (editor). 1994. Rangeland cover types of the United States. Society for Range Management, Denver,
CO. 152pp.
9
Diamond, D. D., and F. E. Smeins. 1984. Remnant grassland vegetation and ecological affinities of the Upper Coastal
Prairie of Texas. Southwestern Naturalist 29:321–334.
10
Diamond, D. D., and T. E. Fulbright. 1990. Contemporary plant communities of upland grasslands of the coastal sand
plain, Texas. Southwestern Naturalist 35:385–392.
11
Chapman,
D. C., D. M. Papoulias, and C. P. Onuf. Not dated. Environmental change in South Texas. http://biology.
usgs.gov/s+t/SNT/index.htm. 11pp.
13

14
Notes

Lower Laguna Madre, Wilacy Co., Texas
WATER
15

WATER
South Texas Moisture... or Lack Thereof
Much of South Texas can be described as arid land occasionally punctuated by floods caused by
hurricanes or Gulf Coast moisture. Rainfall patterns are unpredictable, and drought cycles can last
for several years with rainfall averages as low as several inches per year. Under the most normal
circumstances in South Texas, two rainfall peaks occur during May/June and September/October.
Planning for water needs under these conditions is anything but easy. Water demands by municipalities,
industries, and agricultural irrigators continue to escalate as our population increases. The
amount of water needed in rivers, streams, and coastal bays to support fish and wildlife habitat is
an important issue to resource managers in Texas. Many local economies depend on these flows
to provide income which may be sustained through fishing, hunting, and tourism.
Irrigation
Water quality is important to consider when choosing a water source for irrigation, livestock, or wildlife.
Irrigation water can contain essential plant nutrients; however, certain nutrients can be toxic
to some plants if levels are too high. Water suitable for drinking may not always be acceptable for
growing plants. Appropriate tests should be conducted before deciding upon a water source.
Water quality is based on physical, biological, and chemical properties. Physical properties include
suspended solids such as soil particles, and temperature. Biological properties include algae,
microbes, and disease organisms. Algae can clog irrigation systems, and microbes and disease
organisms can harm wildlife and livestock. Chemical properties are the most critical parameter for
irrigation water, and include soluble salts, hardness, sodium and chloride concentration, and pH.
Iron and boron are also important to consider; imbalances can tie up nutrients and cause chlorosis
to growing plants. Boron is an essential element at low concentrations, but becomes a soil sterilant
at moderate to high concentrations. The most common irrigation water quality problem in South
Texas is high salt and sodium levels, which is also referred to as “saline” water. This condition
causes interference with magnesium and calcium availability to plants. Total salts can be measured
by monitoring the electrical conductivity of the solution; a simple test to determine sodium
adsorption ratio (SAR) can be conducted through most water and soil testing laboratories.
Water quantity and distribution can dictate grazing patterns and distribution of livestock and wildlife.
Over time, these patterns can affect the long-term management objectives of your restoration
project. It is important to distribute water across the landscape, or use water to regulate grazing
pressure and timing.
American alligator (Alligator mississippienis)
17

18
Major Aquifers in Texas

Minor Aquifers in Texas
19

Planning Around Water
• Erosion can be a formidable enemy to restoration efforts. An adjacent watershed’s water quality
can be impacted by runoff caused by eroding bare ground. Be sure to plan for erosion control
and incorporate soil stabilizing techniques or products into your project.
• Include timing of rainfall into project planting schedules.
• Manage livestock and wildlife watering facilities to control animal use in newly restored areas.
• Before drilling water wells, consider availability of underground water and water quality for your
specific purpose.
It is critical to gather information pertaining to water issues in your area, such as
historical rainfall, data evaporation losses, groundwater quality and quantity, river
and stream watershed runoff, and regulations for drilling water wells and constructing
and registering water impoundments, before initiating habitat restoration.
USEFUL REFERENCES
Groundwater and watershed information, drilling regulations, and well water depths and water quality:
Texas Water Development Board
1700 North Congress Avenue
P.O. Box 13231
Austin, TX 78711-3231
512-463-7847
Fax: 512-475-2053
http://rio.twdb.state.tx.us/home/index.asp
Local Water Well Drilling Companies
Rainfall Information
http://www.climatesource.com
http://www.noaa.gov/
Conservation and Grazing Issues
Natural Resource Conservation Service
http://www.tx.nrcs.usda.gov
Other general sources:
Texas Commission on Environmental Quality
San Antonio 210/490-3096
Corpus Christi 361/825-3100
Harlingen 956-425-6010
Laredo 956-791-6611
http://www.tceq.state.tx.us/
Texas State Soil and Water Conservation Board
254-773-2250
http://www.tsswcb.state.tx.us/
Texas Natural Resources Information System
512-463-8337
http://www.tnris.state.tx.us/
20

Pila in Live Oak Woodland, Kenedy Co., Texas
21

22
Notes

Wildflowers near Hebronville, Jim Hogg Co., Texas
PLANNING
YOUR PROJECT23

PLANNING YOUR PROJECT
The success or failure of any restoration project will be directly
proportional to the quality of the initial plan. Time spent carefully
considering details beforehand will be the best investment you can make.
The following outline presents steps for successful completion of a revegetation
project. The sequence will depend on the purpose of the project and current condition
of the site to be re-vegetated.
Define Your Goal
Step 1: Determine your objectives
• Will area be used for activities such as wildlife, livestock, or recreation?
• What is your expected time line?
Step 2: Determine political or legal constraints to the project
• Ensure that appropriate public agencies have been contacted and regulatory requirements met.
• Contact neighboring property owners if they will be affected.
Step 3: Secure maps and aerial surveys of the site, if possible
• The United States Department of Agriculture - Natural
Resource Conservation (USDA-NRCS) Soil Surveys for each
county include critical information on soil types, vegetation
communities, and individual plant lists. They can be accessed
at http://websoilsurvey.nrcs.usda.gov.
• Ecological Site Descriptions are currently being developed by
the USDA-NRCS for each county; drafts are available at http://
esis.sc.egov.usda.gov. A CD-ROM should also be available in
the near future.
The most limiting
factor for all restoration
sites in South Texas is
seed availability. It is
important to work with
local ecotypes
if possible.
• Topographical maps are available through the U.S. Geological Survey: http://
www.tnris.state.tx.us
• Private aerial photography services offer unique options that can be tailored to
the objectives of your restoration project.
25

Step 4: Assessment
• Determine historical vegetation of the site
– This may be difficult to determine, but if local remnant vegetation patches exist in the area,
try to model your restoration efforts after the species composition and distribution of these
sites.
– Visit the site during several seaons of the year; this will allow you to see the variety of plants
(e.g. annuals) that appear at different times.
• Refer to the USDA-NRCS Ecological Site Descriptions for a list of plant species in your area.
• Assess the physical and biological characteristics of the site.
– Identify and characterize soils
– What is the soil texture? Fertility? Salinity levels?
– Evaluate topography
– Is the site rocky, steep, or in a floodplain?
– Determine whether land use on surrounding boundaries will affect your site, such as erosion
caused by oil and gas activity or off-road vehicle trespassing, and how you will correct this.
– What are impacts from grazing/browsing animals (cattle, deer, rabbits, etc.)?
– Determine present vegetation;
– Determine if native plants exist on the site
YES, native plant species exist on the site:
• Identify native plant species on site;
• Identify plants to be salvaged or harvested;
• Salvage and harvest seed;
• Salvage topsoil and subsoil, if appropriate;
• Replace salvaged topsoil;
• Eradicate/control weeds if present (a weed is a plant that interferes with
management objectives for a given area of land at a given point in time);
• Prepare seedbed, amend soils;
• Replant salvaged plants or seeds, or purchase commercial seed adapted to
your area, ensuring that plants are suitable for the site. (shade, soil, water
requirements);
• Control erosion, and mulch (if practical);
• Maintain and monitor (proper grazing management, weed control).
HELPFUL TIP
Many areas of South Texas have soils depleted of nutrients and organic matter, or are missing the upper
layer of topsoil. It may be necessary to adjust the selected species under these conditions. Topsoil and
subsoil is commonly salvaged on mining, and oil and gas drilling locations.
26

NO, native plant species do not exist on the site:
• Conduct brush control or removal of non-native species if necessary;
• Select species for seeding/transplanting;
• Select planting techniques;
• Determine plant/seed sources (purchase commercial seed/plants adapted to
your area, or contract to obtain harvested wild seed/plants);
• Control weeds if present;
• Prepare seedbed, amend soils;
• Determine amount of seed to grow, or number of plants to grow or purchase;
• Control erosion, and mulch (if practical).
HELPFUL TIP
Natural plant populations contain considerable genetic diversity or variation which helps each species
adapt to environmental changes. Plant diversity allows for flexibility and resilience to environmental
disturbances and may result in higher wildlife diversity as well.
Step 5: Decide what species or mixes to plant
• What is the size of the project/area?
• How much seed is needed?
• Is seed available for collection or commercially available for direct seeding or growing
transplants?
• What are the advantages of purchasing vs. harvesting seed?
Step 6: Define your budget and include the following:
• Site preparation costs;
• Planting and harvesting equipment costs;
• Seed and plant material costs;
• Labor costs;
• Annual maintenance costs, such as prescribed fire,
and chemical and mechanical weed suppression;
• Find out if technical assistance and state, federal,
or private funding is available for your project;
• USDA-NRCS offers funding and technical
assistance through a variety of programs:
– http://www.nrcs.usda.gov/programs/
• Texas Parks and Wildlife Department provides technical assistance through the
Private Landowners Managing Natural Resources (PLMNR) and the Landowner
Incentive Program (LIP):
– http://www.tpwd.state.tx.us/landwater/land/private/lip/
HELPFUL TIP
Climate, soil,
existing vegetation, and landscape
features are all important factors
when considering any restoration
project. Remember, more variation in
these features will likely
require more variation
in your species!
At the writing of this handbook, very few species of native ecotypes of South Texas are commercially
available. Check with suppliers in the Index along with your local NRCS representatives to find out about
commercial seed availability.
27

Step 7: Implement your schedule
• Allow 18 to 24 months to plan a restoration project and ensure availability of seed and plant
materials;
• Your project plan should include a detailed schedule of when you will secure the plants and seed
used for the restoration effort;
• Consider whether you have sufficient growing time, and how temperatures affect establishing
plants;
• Prepare the site according to the planting time, and in synchronization with rainfall patterns;
• Consider delaying implementation if the site is currently under drought conditions.
Step 8: Site maintenance
• Maintain and monitor proper grazing management;
• Monitor and continue weed control;
• Apply appropriate management strategies (i.e., prescribed fire);
• Minimize soil disturbances.
HELPFUL TIP
When to Consult a Professional
The following sections will provide general guidelines for carrying out your project, however, this guide
cannot include detailed information for every scenario. For large projects, or for those in particularly
challenging environments, we advise working with experts. You may choose to work with a local resource
agency or a private consultant. In either case, you should be as informed as possible about techniques
and plant species used in your project.
28
Hogplum (Colubrina texensis)

Shrubby oxalis (Oxalis berlandieri)
29

30
Notes

Wildflowers near Laredo, Webb Co., Texas
METHODS OF ESTABLISHING
NATIVE 31
PLANTS

METHODS OF ESTABLISHING
NATIVE PLANTS
There are many different methods used to establish native plants. The method used depends
largely on the availability of seed and other plant materials adapted for the site. Different methods
can be combined to maximize the benefits of each approach to increase the success of the overall
restoration project. Selecting a method suitable to your site is one of the most important considerations
when planning your restoration project.
Cultivated Native Seed
Planting cultivated native seed is a feasible
and practical approach for large-scale
projects when locally adapted seed is
available. Cultivated native seed supplies
are sold commercially and are usually
available in large quantities. This seed is
grown under conditions that minimize the
effects of environmental factors that can
negatively affect seed quality in wild plant
communities. This provides consumers with
a more uniform and consistent product. Most
cultivated seed is tested by independent seed
labs that provide seed quality information in
the form of seed tags or labels.
Cultivated hooded windmillgrass at the USDA-NRCS Plant Materials
Center, Kingsville, TX
These data provide the consumer with the
necessary information to calculate more accurate seeding rates. Cultivated seed can also be
cleaned with machinery to remove non-seed material and weed seed. Seed with fewer contaminants
has higher purity and is easier to plant with conventional machinery.
Cultivated native plants may or may not be selected to produce plants adapted to particular
environmental conditions or with specific characteristics such as higher forage quality. Most
cultivated native seed is sold as a single species. Cultivated seed is a suitable option for those
interested in establishment of native species that are available commercially, or for
those interested in supplementing a native seed harvest.
Planting cultivated native seed is particularly effective for low growing species that
might be missed when harvesting from a wild plant community. Desirable species
not present during time of wild harvest can be purchased and added to the wild mix.
Obtaining permission to harvest native wild seed on private lands can be complicated
and unpredictable. A benefit of cultivated commercially grown seed is its availability
and accessibility to the public.
33

Wild Native Seed
Indian blanket (Gaillardia pulchella)
Native seed can be harvested from wild stands dominated by a desired
target species or from stands with a mixture of species. Harvesting wild
native seed is a good method for restoration when donor sites are available
that have similar environmental conditions to the restoration site. Important
environmental conditions to consider when comparing sites are soils, rainfall,
temperature patterns, and topography. This method of harvesting seeds for
restoration should also be considered if similarly adapted commercial native seed is
unavailable, or when seed costs are prohibitive.
Because native wild populations are so dynamic, the
abundance and diversity of species at a particular site
will vary with variation in environmental factors.
Species can be harvested individually or as a mixture,
either by hand or with mechanical seed harvesters.
Wild native seed should be harvested when seeds of
the majority of the target species have matured and
are ripe. No particular harvest technique or collection
period, however, will provide seeds from all of the
species in a site because different species flower
at different times of the year. Several harvests are
therefore recommended throughout the year if the
Collecting seed with a hand-held seed stripper
objective is
to maximize the number of species in the mix. Harvesting
during multiple years may increase the number of species
in the mix because certain species may produce seeds
only in wetter years. Typically, wild harvests are not
cleaned as thoroughly as cultivated seed because they
vary greatly in size, purity, and maturity. Seeding rates
of native seed mixes are difficult to determine because
of the diversity of species and the variable quantity and
quality of the seed harvested.
Collecting seed by hand
34
The high quantities of non-seed material in wild
native seeds make application with mechanical
equipment such as rangeland seed drills or
broadcasters difficult. For an additional cost, wild
harvested native seed can be sent to commercial
seed companies for cleaning to remove nonseed
material, resulting in less bulk for storage
and a cleaner mix for planting. Procedures for
certification have not been developed to test
native seed mixes.
find new photo
Harvesting wild seed

Wild Native Hay
Harvested wild native hay can be used to re-establish native grasses and forbs. Planting with wild
harvested native hay is especially useful on rough, erosive terrain. The hay serves as the seed
source and provides some protection from wind and water erosion. Organic material in harvested
hay increases soil organic matter and may create a more favorable environment for soil organisms.
The first step is to locate donor sites that have suitable terrain for mechanical harvest.
Environmental conditions should
be similar between the restoration
site and the donor site. This site
should have livestock removed
well in advance of the cutting to
allow the plants to set seed. The
area to be hayed may be cut at
various times throughout the
year if rainfall is adequate. This
should increase species diversity
of the collected seed because
seeds of different native species
mature at different times of the
year. Hay should be harvested
when seed is ripe but before
excessive shattering occurs.
Not all species mature at once,
so focus on dominant species or
on the species you desire most
to plant.
Spreading native hay
Natural Revegetation
Natural revegetation of native species is an option for degraded areas with a sufficient native seed
bank in the soil. This method focuses on the repair of ecosystem functions and requires good
understanding of individual components of plant community succession, and how to monitor and
fine-tune management practices. Disadvantages include the extended period of time that natural
revegetation requires, and the possible invasion of non-native species before establishment of
native plant communities.
False rhodesgrass (Trichloris crinita) and plains bristlegrass (Setaria leucopila)
35

Transplanting
Transplanting native plants is
another option for revegetating
degraded habitats. This method is
used in very small areas or areas
in need of rapid establishment
of native vegetation. This could
include areas prone to significant
soil erosion or to areas with high
probability of non-native plant
invasion. Transplanting native
plants is costly, and not feasible
for large revegetation projects.
Transplants can be moved
from native sites, or grown in
Transplanting seedlings
containers. Transplants can
quickly establish in new areas
because of established root
Transplanting grass seedlings
systems. This method is also
good to use when a desired target species is difficult to establish by seed, has poor seed production,
and/or is considered sensitive or rare.
Integrating Methods
Cultivated native seed, wild native seed, wild native hay, or transplants can be integrated to
achieve the goals for your restoration project. A combination of methods suitable to your site
may increase the probability of success, and add more diversity to the restored community. For
example, cultivated native seed and wild native seed may be planted together. Wild seed may
increase species diversity and cultivated seed may give the volume of seed needed to plant the
total acreage.
Bristle leaf dogweed (Thymophylla tenuiloba)
36

Common sunflower (Helianthus annuus)
37

38
Notes

Mudflats adjacent to Lower Laguna Madre, Cameron Co., Texas
SOILS
39

SOILS
Soils play a crucial role in the restoration process. Physical and chemical properties of soils,
including the limitations imposed by them, must be considered in restoration efforts. Evaluating
your soils thoroughly before implementing a restoration project is important for site preparation
and planting strategies. Resources are available that will help you recognize soil characteristics
specific to your restoration site. Soil surveys for most South Texas counties are available from
the USDA-NRCS (Appendix F). These surveys will be a starting point for general information on
soil types, soil texture, factors affecting limitations in the soil, and land use practices. Local NRCS
specialists can also provide soils information.
Soils have chemical, physical, and biological characteristics that constantly interact with plants.
Plants, soil, and microbes form complex ecosystems resulting in interactions that can benefit or
harm native plants. The key is to find a balance that optimizes native plant productivity and survival,
and ultimately results in restoration. Unfortunately, little information is available about this topic,
and experimentation may be required.
Importance of Soils to the Restoration Process
Soil is the environment where the biological and mineral worlds interact. It serves as the medium for
plant growth, and is comprised of minerals, dead organic matter, and living organisms such as soil
microbes, fungi, worms, insects, and roots. Soil can also contain seed, which is the basis for plant
reproduction and long-term survival of plant communities. All of these characteristics influence
overall plant community composition, and are important to consider when selecting species for
planting. Species adapted to local soils will perform better than species that are not.
Re-establishing native plant populations often involves more than just replanting or
reseeding vegetation. Changes in land use can directly impact soil, and can cause
degradation to the point where the soil no longer supports plant growth. Invasive,
weedy plants may out-compete natives in overly disturbed and fertilized soils.
An understanding of soil processes, physical and chemical properties, and
historical land use practices are extremely important for successful
re-establishment of native plants.
Physical Properties
Soils have many physical properties. Soil profiles
are made up of horizons, or layers of different soils.
Typical soil profiles are made up of 3 general layers:
topsoil, subsoil, and parent material.
41

The upper layers, or horizons, of a soil profile contain significantly higher amounts of organic matter
than the underlying layers because most biological activity occurs in the uppermost horizons. Soils
are classified by a combination of soil profiles and a number of other physical properties. Some of
these properties include structure, surface texture, depth, slope, and permeability.
Soil structure refers to the relative proportions of air and solid matter in the soil as well as the size,
shape, and arrangement of these aggregations. Soils are not just collections of loose soil particles;
they are composed of clods, clumps, and chunks of sand, silt, and clay particles mixed with organic
material. Soil scientists use categories of structure to describe these aggregates, such as bulk
density, which is the amount of solid material per unit volume (g/cm 3 ) in the soil. A soil that provides
ideal growing conditions contains about 50% solid volume and 50% pore volume. A high bulk
density translates to a greater degree of soil compaction and less developed structure.
Soil surface texture
is the primary factor
to consider when
planning a restoration
project. Soil texture
is determined by the
size of the particles
that make up the soil.
Particles are divided
into 3 size categories:
sand, silt, and clay.
Sand is the largest
particle; clay is the
smallest. Soil types
are based on the ratio
of these particle sizes
within the soil. Soil
texture influences
how quickly water,
nutrients, and oxygen
infiltrate and percolate
through the soil, as
well as the quantity of
water and nutrients
a soil can retain. In
general, soils high in
clay do not drain well.
Soil textural triangle
They retain nutrients but have a tendency to become waterlogged. Sandy soils drain well, but have
low nutrient-holding capacity. Loamy soils represent a balance between sand and clay, and are
preferred for agricultural use because they retain nutrients and drain well. Soil surface texture also
has a significant influence on the types and species of native plants that can grow there. Accurate
determination of soil texture should be a major factor in deciding which plants to use for establishing
vegetation on a restoration site. Plants that are poorly adapted to a particular soil texture will most
likely be unable to persist there.
42

Soil depth and slope of the site can play an important role in determining how to prepare a restoration
site for planting. Shallow soils or those susceptible to erosion must be carefully prepared and
maintained in order to minimize soil loss. These sensitive areas should not be left without some
type of cover to hold soil in place. For areas prone to significant erosion, restoration plans should
include soil conservation practices such as the use of cover crops, mulches, or non-conventional
tillage and planting methods.
Soil permeability refers to the ability of water and nutrients to permeate throughout a soil horizon.
Soils with too low or high a degree of permeability can reduce the ability of a site for plant growth.
Soil permeability can be affected by factors such as slope of the site, soil texture, and compaction
of the soil. Soil compaction is the compression of soil particles, which can be caused by continual
traffic (animal or human) or heavy machinery. As soil compacts, particles move closer together
creating fewer air spaces, and prevent water and roots from moving through the soil. Furthermore,
seeds cannot establish in compacted soils, but instead remain on top of the soil and become
vulnerable to the elements and seed-eaters such as birds and insects.
Organic Matter in Soil
Organic matter is composed of decaying plant material mixed with newly synthesized compounds
formed by microbes. It is also the main source of nutrients that supports microbial populations,
insects, and worms, which in turn, support vigorous plant growth.
Organic matter serves a variety of purposes in soil. As organic matter breaks down, plant nutrients
such as nitrogen and phosphorus are released for root uptake. Poorly performing native plantings
are often associated with a lack of organic matter in the soil. Organic matter improves soil structure
by making it less susceptible to erosion, and helps to increase water infiltration and retention in the
soil. For example, adding organic matter to sandy soil can improve its nutrient and water holding
capacity, and may actually be the main factor for water retention. If added to a clay soil, organic
matter can improve water infiltration and percolation. The long-term accumulation of organic
matter is also very important, and can be ensured through proper
rangeland management practices. Overall, an appropriate
amount of organic matter will result in higher plant survival.
Frequent cultivation results in rapid depletion of soil organic
matter in South Texas.
Soil Microbial Activity
Microbial activity plays an important role in
establishment and persistence of native plants. While
plants get nutrients directly from soil, other nutrients
are made available to plant roots through microbial
activity. Organic matter is the key to microbial activity
in the soil. Nitrogen, for example, is supplied mainly
through microbial decomposition of organic matter. In
most cases, active microbes in the root zone of the plant, or
the rhizosphere, help increase plant nutrient uptake.
43

Another example is phosphorus, which is required in relatively large amounts by most plants.
Some phosphorus is absorbed from soil minerals and organic matter directly through the roots, but
root-inhabiting microbes, especially mycorrhizal fungi, enhance availability. Mycorrhizal fungi are
symbiotic with nearly all plants, and can be particularly important to native plants. These nutritional
interactions are often ignored in commercial agriculture, but they are vital to plant survival on native
landscapes. A healthy diverse microbial community in the soil greatly increases plant productivity.
Microbial species richness and diversity are important because they allow for a more varied and
flexible response to envinronmental fluctuations and stress. Soils with high species microbial
diversity are more likely to cope with disturbances or drought.
Chemical Properties
Several chemical properties of soil should be considered before a restoration project is implemented.
These are pH, electrical conductivity (EC), sodium absorption ratio, cation exchange capacity
(CEC), and percent organic matter. These chemical properties are important because they
greatly influence the suitability of soil for seed germination, plant survival, and growth. The only
way to determine the chemical properties of a soil is to have it tested.
Topsoil
Topsoil is the most critical soil layer when it comes to native plant restoration. It is defined as the
uppermost 6 to 12” surface layer of undisturbed soils, where the bulk of the root zone is located.
This portion of the soil contains large reserves of plant nutrients and the organisms that influence
soil nutrient cycling, and is a critical component to a healthy ecosystem. Biological activity is
usually limited below the topsoil layer.
The advantages of using natural productive topsoil in restoration
include the presence of organic matter, microbial populations,
and native seed that are invaluable to the re-vegetation
process and cost effective to the manager. When topsoil has
eroded or been removed from an area, it may be possible to
purchase native topsoil from areas adjacent to the restoration
site, when feasible. In the event of severely damaged sites, such
as with oil drilling pads, it may be necessary to conduct a soil test that will
reliably determine the quality of the topsoil. If inadequate, a commercial grade
One of the
greatest limiting factors
in restoring native
vegetation is the loss of
topsoil.
of topsoil can be purchased in bags or in bulk. Bagged topsoil is usually sold in 40
to 50 lb quantities and has been amended with lime, fertilizer, and organic matter.
Bulk topsoil generally is a native soil taken from the surface and sold in truckload
lots. In many cases, modifying soil properties may be necessary where native soils
have been damaged. Soils can be amended with a range of materials that counteract
problems contributing to the loss of native plants from the site. Soil amendments also
improve water use efficiency, increasing plant survival.
44

WHAT IS HUMUS?
A foundational part of organic matter is humus. Humus is a chemically complex mixture of brown
to dark colored organic substances that form during decomposition of plant and animal residues,
and makes an excellent substrate for plant growth. In addition, carbon compounds contained in the
residues that were synthesized by the plant or animal when it was alive become protein and energy for
various bacteria, protists, fungi, and actinomycetes.
Aerobic microorganisms (requiring oxygen) are most adept at breaking down organic matter, so the amount
of oxygen in the soil is important. Soil moisture, temperature, and carbon:nitrogen ratios are also factors
in the rate of decomposition. Anaerobic microorganisms (not requiring oxygen) also break down organic
matter (such as that found in water), but at a much slower rate. In the long run though, they can
produce more humus (called “muck” or organic soil). In soils that contain too much oxygen (sand, for
example), very little humus is formed.
Soil Amendments
Most of what we know about soil amendments originates from the agricultural industry, so caution
should be used when applying this knowledge to native plant habitats. Differences between the
way native and crop plants respond to varying soil problems should be thoroughly considered.
For instance, adding chemical fertilizers to increase soil fertility and boost plant productivity in
agriculture is a common practice. Native plants, however, may be less
responsive to increased fertility than invasive or weedy species, and might
be out-competed and displaced by weeds in a heavily fertilized soil. This
often occurs in previously cultivated soils that contain residual fertilizer.
Soil pH
Soil pH has a strong effect on nutrient availability to plants. Maintaining a near
neutral pH usually benefits trace element uptake such as with iron, zinc, and copper.
Because most soils in South Texas have an elevated pH, materials such as sulphur,
or acidulants, chemically react in the soil to produce acids. When elemental sulfur
oxidizes through microbial action, sulfuric acid is formed. This reduces the pH and
provides the plant with sulfate, which is often deficient in soils. Acid-forming fertilizers
such as ammonium sulfate also serve a similar purpose; the impact of acid additions on
most soils in South Texas, however, is minimal because of the difficulty in neutralizing
such large amounts of calcium in the soils. Limestone and dolomite (a mixture of
calcium and magnesium carbonates) are often used to increase soil pH.
Remember,
pH < 7.0 is acidic
pH = 7.0 is neutral
pH > 7.0 is basic
52
MICROBES
Microbes are always closely associated with organic materials, and can occupy particular niches in the soil
if they are available. Microbes are generally separated into 6 functional groups: aerobic, anaerobic, and
nitrogen fixing bacteria; actinomycetes, pseudomonades; and yeasts and molds (fungi). Bacteria act as
decomposers, helping to provide important nutrients to plants. For example, the Rhizobium bacterium is
symbiotic with legumes, which “fix” nitrogen from the air and makes it available to other plants. Actinomycetes
break down lignin and cellulose in the soil, such as insect remains, and can survive and function at high
temperatures. Much of the heat associated with decaying matter in soils is a result of actinomycetes. Fungi
help to decompose organic matter in the soil, and help bind soil particles together. Mycorrhizal fungi, for
example, are symbiotic with growing roots and improve plant absorption of phosphorus, iron, and water.
45

THE EFFECTS OF LAND USE PRACTICES ON SOIL
Thousands of acres of rangeland in South Texas have been manipulated with numerous mechanical methods that
have negatively impacted soil properties. Root plowing, for example, is commonly used for brush removal. In areas
of South Texas where the topsoil layer is thin, root plowing can rearrange the soil profile by burying the A horizon,
which contains the greatest percentage of the seed bank, and bringing the B and C horizons to the surface. The B
and C horizons contain higher clay content and often produce a hard, compacted surface or “hard pan,” making it
difficult for water to penetrate. Because mechanical treatments alter soil properties, historical mechanical alteration
of the habitat should be taken into consideration before implementing restoration efforts.
Overgrazing has also affected South Texas soils. Continuous overuse of rangeland vegetation by grazing and
browsing animals has led to loss of organic matter, increased soil compaction, and the loss of the A horizon on many
areas. Long-term absence of vegetation caused by heavy grazing coupled with drought has also led to erosion
problems in some portions of South Texas. Areas that have undergone extensive erosion and loss of topsoil may
require significant soil rebuilding efforts in order to implement restoration strategies. Consult your local NRCS
specialist for more information regarding erosion control and management.
Farming practices have also greatly influenced soils in some portions of South Texas. Soils that have been used for
farming should be carefully evaluated and amended to support native revegetation efforts.
Introduction of non-native grasses and spread of noxious brush species have also detrimentally affected productivity
of soils in our region. For example, non-native grass seed present in the soil can readily sprout and, as a result,
reinvade an area even after removal of problem plants. Once a non-native plant becomes established, the soil likely
becomes contaminated with undesirable seed. Some non-native species have proven difficult to control, and are
efficient at out-competing native species.
Water Infiltration, Sodium, and Salinity
Water infiltration problems are common in soils with high sodium or clay content. High sodium also
causes osmotic problems for plants, affecting water use and nutrient flow. Soils high in sodium are
often salty and exhibit crusting problems that keep water from penetrating into the soil, resulting
in erosion. Gypsum has often been used to leach sodium from the soil profile, helping to build up
the soil structure and improve infiltration. Liquid calcium is frequently used to leach sodium from
the upper layer of soil to aid vegetation establishment on brine spills. Organic amendments such
as composts, animal manures, and sludges are also used to stimulate soil microbes to produce
polysaccharides and other organic compounds that bind soil particles into larger clumps. These
aggregates have larger spaces between them to allow water to infiltrate more freely into the soil.
Synthetic materials, such as polyacrylamide, can also be used to promote aggregation, but may not
be economical or produce consistent results.
Water Holding and Cation Exchange Capacity
Water holding capacity (WHC) is a particle surface property. The greater the particle surface
area, the larger the amount of water can be held in the soil. Sandy soils have low particle surface
areas and thus low WHC. Clay soils, on the other hand, have high particle surface areas and a
high WHC. Cation exchange capacity (CEC) is the ability of a soil to hold cation nutrients. CEC
follows the same general principle; sandy soils have low CEC’s and clayey soils have high CEC’s.
Both WHC and CEC can be improved by adding organic amendments to the soil. Composts with
a high content of humus improve WHC and CEC, and provide the plant with fertilizer and trace
elements. Zeolites and diatomaceous earth are naturally occurring inorganic amendments that can
also improve WHC and CEC by increasing long-term water and nutrient holding capacities.
46

HELPFUL TIPS
When is the Best Time to Salvage Topsoil?
Topsoil should be salvaged when moist, but not wet. Finely textured soils can be damaged if salvaged when they are
too wet; alternately, if soil is too dry, important living plant material could be lost 1 .
Storing and Protecting Salvaged Topsoil:
Topsoils should only be stored for brief periods of time to ensure that important microbial processes are not interrupted.
If soil must be stored more than 6 months, protecting the soil with a cover crop may be an option.
Allow sufficient exposure to air; store soil in shallow piles, preferably less than 3 feet.
Keep soil isolated from invasive weeds or non-native plant species.
Topsoil Application:
• Apply topsoil in an uneven pattern, avoiding too much manipulation; this imitates natural patterns with variable soil
depths.
• Avoid excessive machine passes to prevent compaction.
• Apply topsoil within a few days of seeding; this will limit invasion of weedy species, and soil loss caused by
erosion.
• If topsoil is imported from another location, carefully evaluate the soil to ensure that invasive or non-native plant
species are not being introduced to the site.
Increasing Fertility with Organic Amendments
A variety of organic materials are available for land application, the majority being raw wood wastes,
composts, animal manures, and sludges. With the exception of wood wastes, these amendments
are fairly rich in plant nutrients. As waste decomposition slowly takes place, nutrients are gradually
metered out to young plants. In the hot, dry environment of South Texas, building organic matter
is challenging because of the rapid oxidation; growing cover crops, however, is an efficient way to
add organic matter to soil.
Organic matter amendments may be the best way to remedy several soil problems at once, but too
much organic material can make the soil excessively rich and allow weeds a growth advantage.
Adding the organic amendment to a small area immediately around the native plants, either in a
row or individually, may help minimize the weed problem. This will stimulate the native plants’
growth, but keep the remaining soil less fertile and less friendly to invasive weeds.
Some organic amendments, such as wood wastes and sawdust, can tie up nutrients as they
decompose in the soil. Amendments with high carbon-to-nitrogen ratios commonly cause this
problem. While the nutrient tie-up is temporary, the deficiency may still contribute to a planting
failure. It may be necessary to supplement nutrient-deficient organic amendments with fertilizer
elements to counteract this problem.
Adequate levels of organic matter are usually an indicator of the presence of microbes, but
sometimes organic amendments are not enough to create a healthy growing environment for newly
planted native plants. Inoculation of beneficial microbes may be necessary to enhance native plant
success, and to restore local soil communities. Commercial inocula of soil microbes are available,
and can be beneficial in re-establishing native plants. It should be noted, however, that some
species in commercially available inocula are not indigenous, and may contradict goals to maintain
a native system. Efforts should be made to obtain native and locally adapted microbes.
47

There are several approaches to microbial inoculation. Pelleting native seed with mycorrhizal
fungi, other microbes, and nutrients may supply much of what native plants need for establishment.
Researchers are attempting to further understand the full potential of these treatments, but current
findings demonstrate that for selected species like legumes, they can greatly improve the success of
native plant restoration. Microbial inoculation can be accomplished by incorporating small amounts
of newly harvested topsoil from a nearby area into the existing layer of soil in soils that have been
greatly degraded or where topsoil has been removed. Use of compost tea, a liquid produced by
leaching soluble nutrients and extracting microorganisms from compost, is another common way
to increase microbial populations.
Biological Problems
Soil amendments can sometimes be expensive or impractical, especially for large-scale projects.
Where they are used, however, the long-term benefits they afford to plants can make a difference,
and can greatly improve the chances that plant populations will recover, and ultimately re-establish.
Proper rangeland management, however, can be the most profitable means of enhancing soils and
should not be overlooked when developing soil improvement strategies.
SOIL TESTING
Soil tests are reasonably priced and worth the valuable information obtained from test results. A soil test
not only helps in the selection of species best suited for each site, but also identifies potential toxicity
problems. A basic soil test provides information on the 4 soil properties listed earlier, and precise soil
texture readings. Details on the macronutrients (nitrogen, phosphorus, and potassium) and micronutrients
(copper, iron, magnesium, and zinc) present in the soil are also provided. In some cases, further testing
may be necessary for measuring the quantities of salts and heavy metals, as well as other physical
properties, especially if high salinity levels or contaminants such as hydrocarbons exist.
Some laboratories offer assistance with interpreting test results. Recommendations given for soil
amendments are usually based on agricultural standards, however, and may not always be applicable to
native plant growth. Restoration professionals can give you advice on how to apply these results.
48
Soil testing in the field

COLLECTION OF SOIL SAMPLES FOR TESTING
Items Needed for Soil Sampling
1. Map of area (aerial topographic or USDA-NRCS Soil Survey for your county);
2. Soil punch, sharpshooter, or trowel;
3. Plastic freezer bags;
4. Permanent marker; and
5. GPS unit (optional) for future reference to site.
How To Collect the Samples
Most testing facilities provide specific instructions regarding procedures for collecting soil samples.
Following are some general guidelines:
1. Collect several subsamples (4 to 20) from each distinct soil type in an area using a consistent
sampling method that ensures a uniform representative sample of the area. Mix subsamples
together to form a composite for each soil type. Always take a minimum of 4 samples to
have a representative composite, even for small areas.
2. Take a sample from each composite. Remove rocks and roots, if present.
3. Label the sample with date and location of collection.
4. If you suspect a problem area, such as a contaminant or high levels of salts, keep the sample
separate. To sample contaminated soils where it may be necessary to use the test results
for further action, contact a professional testing lab to ensure proper procedures are followed
for handling and collecting the samples.
5. Send the soil samples to the laboratory as soon as possible after collection to ensure accurate
results.
You may make several composite samples representing different soil depths, but at least one composite
sample should be taken from the soil surface, at a depth of 0-6”. For small areas, testing one composite
soil sample will be sufficient. For larger areas, several composite soil samples may be necessary,
particularly if variation in soil color/texture is noted. If you can see a difference, the soil will test differently.
Samples may be removed with a soil corer or a shovel, as long as the sampling method is consistent and
the tool is not contaminated with soil from another site. Avoid collecting the surface litter layer; otherwise
it may bias the reported organic content for the soil. Composite soils should be as similar as possible;
if obvious soil differences are ignored, analysis of a composite sample may misrepresent the true soil
characteristics. See Appendix F for a list of soil testing laboratories.
USEFUL REFERENCES
Sandy loam soil
See Soil Foodweb, Inc.’s website for information on compost teas. http://www.soilfoodweb.com/sfi_html/index.html
LITERATURE CITED
1
Soil
Rehabilitation Guidebook, March 1997. Ministry of Forests, British Columbia, Canada. http://www.for.gov.bc.ca/
tasb/legsregs/fpc/FPCGUIDE/soilreha/REHABTOC.HTM
49

50
Notes

Partridge pea (Chamaecrista fasciculata)
SITE PREPARATION
51

SITE PREPARATION
Site preparation before seeding or transplanting is very important. Clearing the seedbed will
prepare the site for seedling emergence or transplants, minimizing competition with already existing
introduced or weedy species. This can be done with a variety of chemical or mechanical means,
depending on the conditions present. It is also important to hire or seek the counsel of adequately
trained personnel and range specialists to help advise you with this task. Mistakes can be costly
to both you and the landscape.
Timing is Everything!
Planting dates for restoration projects must be coordinated with the physiological requirements of
germinating seeds, as well as the favorable temperature and moisture conditions of early spring
and fall in South Texas. Planting in dry conditions and hoping for rain is a sure setup for failure
in restoration projects. The majority of South Texas receives peak rainfall in early spring or fall.
Spring plantings should be done early enough to allow plants to establish adequate root systems
before the hot and dry South Texas summers. If planting in fall, plant early enough to avoid freezing
temperatures. Seedlings or transplants must have time to establish adequate root systems in order
to ensure survival throughout winter.
Cropland, or previously cultivated sites, should be prepared and left fallow at least 3 to 6 months
before either of the 2 peak rainfall periods in South Texas. Seeding should take place immediately
preceding these rainfall periods; it is important to allow the soil to absorb as much moisture as
possible to ensure seedling survival.
The site should be prepared before expected peak rainfall periods to restore deteriorated rangelands,
or rangelands that were previously infested with unwanted brush or non-native species. The site
should then be seeded or transplanted immediately after preparation. This will help to minimize
reinvasion of non-native species. Fallow, disturbed sites should not be left uncovered.
Site Preparation Methods
Site preparation techniques vary, depending on soil characteristics, brush density, and whether the
site has been previously cultivated. Good seedbeds can be prepared with a combination of tools
and techniques. Selection of the methods used should be based on the following considerations:
• Does brush removal need to be conducted based on objectives of the project?
• Are weeds and non-native grasses present?
• Is the soil compacted or disturbed to a point that moisture infiltration and retention are
compromised?
• Are soil amendments necessary to facilitate native plant growth?
• Will a cover crop be necessary to prepare the soil in the event native plants aren’t immediately
seeded?
• Is the seedbed well tilled, smooth, and free of large clods?
53

By considering these factors, a site preparation strategy can be developed. Above all, remember
that disturbances to the restoration site should be minimized, and only those techniques that are
necessary to provide an adequate seedbed should be used.
Sites with dense brush canopies or brush densities higher than defined objectives should be
controlled. A variety of brush control methods have been developed for South Texas and consultation
with an expert can be very helpful. Following are basic guidelines for deciding which type of brush
control to use:
Brush Control
Root plowing
Root plowing involves using blades mounted on large
bulldozers that remove the root crown of brush 12”
to 20” below the ground. Subsequently, however,
significant soil disturbance occurs using this
technique. Also, areas that are root plowed
require further mechanical treatments such as
stacking, roller chopping, raking, or prescribed
fire to dispose of uprooted brush; this helps
to prepare the site for planting with seed drills
or transplanting equipment. Unclean root plowed
areas can be seeded aerially or with broadcast seeders,
but effectiveness is compromised by the roughness of the seedbed. Root plowing
Root plow
is the most effective method available to completely remove unwanted brush; brush canopy cover
can be reduced for up to 20 years post-treatment. Root plowing does have some adverse affects,
however, including mixing soil layers and providing an opportunity for weeds and non-native
grasses to establish. Ultimately, this may prevent the establishment of seeded or transplanted
native grasses and forbs, and can result in restoration failure. Root plowing should only be used
in situations where brush removal is necessary because of density, species composition, or when
maturity of brush prevents proper preparation of the seedbed for restoration.
THE BRUSH INVASION
Historical records indicate that much of South Texas was comprised of grasslands previous to
colonial settlement. Several factors contributed to the dense “brush country” that South Texas has
today. Grasslands have evolved under a disturbance pattern that was altered with settlement and
heavy continuous grazing. Fire suppression, coupled with heavy grazing pressure that removed the
fine fuel needed for burning, further altered the cycle. These factors have favored the brush species
found today. Because most brush species respond with “multi-stems” to injury or cutting, brush
removal efforts can prove futile, and brush densities can acutally increase. Recent brush ecology
studies have provided an understanding about brush invasion, and factors that must be considered
in its management.
54

Roller Chopping & Aerating
Roller chopping and aerating are effective tools for the top removal of brush. These practices
also help to alleviate compaction in the upper layer of soil by increasing water percolation and
increasing pore space in soil. Roller chopping and aerating, however, will only suppress, and not
kill, existing brush. These tools should be used in areas that have minor infestations of brush,
Roller chopper
Lawson Aerator
areas in which maintaining brush diversity is important, or in areas where disturbance to wildlife
needs to be minimized. Proper management and timely planting are crucial aspects of using these
treatments; regrowth of brush is inevitable, and subsequent competition with restored plants is a
concern.
Think first!
Brush control can be
beneficial, but if conducted
incorrectly, it can cause more
damage to your site!
Chaining
Chaining involves pulling large anchor chains
between 2 crawler-type tractors across the landscape
and uprooting or disturbing brush. Chaining can be
very effective at controlling mature stands of brush,
because large stems are more brittle and are pulled from the
soil or broken off below the root crown. However, for small or immature
stands of brush, chaining only “prunes” young branches and stems, resulting
in substantial regrowth. In the case of preparing a site for restoration, chaining must
be used in conjunction with other brush control treatments in order to be effective.
55

Herbicides
Brush can also be controlled using herbicides. However, careful consideration should be made
when using herbicides; be sure to consult with herbicide experts about the management objectives
and potential problems involved with this tool. Following are some general guidelines for herbicide
use from the Texas Cooperative Extension:
• Identify the weed or brush species, and evaluate the need for control;
• Consider the expected benefits, costs, and alternative control practices;
• Select and purchase the suggested herbicide for the weed or brush species;
• Provide and require the use of proper safety equipment;
• Calibrate spray equipment;
• Mix herbicides in a ventilated area, preferably outside;
• Spray under conditions that minimize drift to susceptible crops;
• Apply the herbicides at the suggested rate and time;
• Keep a record of the herbicide used, the time required to spray, weather conditions, rate of
herbicide in carrier, date and location, and the person using the herbicide;
• Considering which method of application that is suitable to the site is also important, i.e., aerial,
ground, or individual plant treatment (IPT). Consulting with manufacture representatives
will help you decide which herbicide to use for selected species, and which method will
work best for your situation.
Some common herbicides for controlling South Texas brush are
• Clopyralid
• Hexazinone
• Imazapyr
• Picloram
• Picloram:2,4-D(1:4)
• Tebuthiuron
• Triclopyr
• Triclopyr:2,4-D(1:2)
Weed and Non-Native Grass Removal
The removal of unwanted weeds and non-native grasses is sometimes necessary before restoring
native vegetation. These unwanted plants compete with native restored plants for valuable
resources such as soil nutrients, water, light and space.
Root plowed mesquite and huisache
56

There are several approaches that you can use to control weeds. Conducting several cultivations
before planting will keep well-developed weeds from re-sprouting or re-establishing. Early weed
control during the initial seeding year will also reduce seedling competition for moisture and other
limiting resources. Before seeding, weeds can be controlled with tillage or herbicides. If restoring
cultivated land and weeds are a problem, weeds should be treated before they get taller than 2–4”
both before seeding as well as 6 to 12 months after planting; follow-up treatments will probably be
necessary. In a mixed planting, herbicides are not an option because they are nonselective and will
kill non-targeted plants. Even with ample preparation some weeds will still emerge and compete
with your planted species, so taller weeds may need to be mowed.
Some herbicides that can be used for broadleaf weed control are
• 2,4-D
• Dicamba
• Dicamba:2,4-D(1:3)
• Glyphosate
• Metsulfuron methyl
• Picloram:2,4-D(1:4)
• Tebuthiuron
Non-native grasses can be much more difficult to control. Herbicides and disking can be used to
kill existing plants, but because of the large existing seed bank, these plants will often re-establish.
Because little is known about how to control non-native grasses, experimenting with a variety of
methods may be necessary. As a general rule, however, non-native grasses respond favorably to
increased soil disturbance; minimizing disturbance while preparing a site for restoration might help
reduce competition. When restoring areas infested with non-native grasses, establishing a cover
crop or quickly establishing natives will likely be your best option.
Alleviating Soil Compaction
Soil compaction inhibits water and nutrient cycling; most important, it is detrimental to plants. Soil
compaction is caused by a variety of factors, but mainly the loss of organic material, continual traffic
from humans, animals, or vehicles, or use of heavy machinery. There are a variety of options for
alleviating soil compaction, but roller chopping/aeration (see brush control) and ripping are the
most effective techniques.
Ripping (Subsoiling)
Ripping is used to break up compacted soils so that moisture can be more readily absorbed.
This disturbs soil much deeper than conventional tilling or plowing, and allows more moisture to
accumulate into a soil profile. Ripping is also recommended when restoring farmland into native
habitat; a chisel plow at 8–10” or a paratill at 12–16” are sufficient. Ripping should be followed by
disking to create a uniform seedbed.
SOIL AMENDMENTS
Soil amendments may be necessary to provide a suitable habitat for native plant growth, and
should be added to soils prior to planting.
57

Cover Crops
For disturbed sites where nutrients and organic matter have been depleted, it might be necessary
to improve the soil before establishing native vegetation. In some cases, you may not be able to
immediately plant the native crop. If so, you can protect the soil from erosion by planting a cover
crop. Cover crops can be used to provide an immediate ground cover, and will help build the soil
and start the upward trend in succession. Roots provided by cover crops can also help to stabilize
the soil.
Mulch created by cover crops provides essential nutrients and organic matter, which breaks
down quickly in the hot South Texas climate. Cover crops also help reduce soil temperatures
and evaporation losses. These crops can be annual, biannual, or perennial. When crops are
planted primarily to protect the soil, a mix of legumes and grasses yields the highest cover per
surface area. There are also cool and warm season annual cover crops. In South Texas, most
cool season crops are planted in late fall, and include clovers, vetches, medics, winter peas, and
cereal grains such as oats, tricale, and wheat. Cereal ryes are not recommended because they
can suppress the germination of other plants. Warm season annuals are planted in early spring,
and include cowpeas, soybeans, sorghum, sudangrass, millet, and forage sorghums. Techniques
for establishing cover crops include aerial and ground broadcast applications, and drill seeding.
Seedbed Preparation
The final step in site preparation is to thoroughly prepare the actual seedbed. Each restoration site,
regardless of other treatments, will have to be disked to smooth out the surface before transplanting
or non-aerial seeding; rough seedbeds may reduce the performance of young seedlings and
transplants. Disking should be conducted only deep enough to smooth the surface; this will help
to reduce soil moisture loss.
Mechanical preparation can sometimes result in a loose, fluffy seedbed. This is especially true for
sandy loam type soils; fluffy seed beds are most detrimental to very small native seeds that can be
buried when the soil settles and hardens. To reduce fluffiness of the soil, allow some time to pass
between the last cultivation and seeding. A good rain after cultivation will help in natural firming of
the seedbed.
Cover crop in South Texas
58

Arizona cottontop (Digitaria californica)
59

60
Notes

Harvesting multiflowered false rhodesgrass (Chloris pluriflora)
ACQUIRING SEED
61

ACQUIRING SEED
There are several options for acquiring seed for your restoration project. Locally adapted ecotype
seed releases are available in limited numbers at present, so finding the right quantities and kinds of
seed for your project may be challenging. Nevertheless, within a few years, locally adapted ecotypes
should be readily available on the commercial market. For now, here are some suggestions to help
you get started.
Purchasing Seed from Dealers
After you determine which species you will be using for your project, but before contacting a seed
dealer, it is important to know the following:
• Common and scientific name of the species you are purchasing;
• The particular species and variety of that plant best suited for your
restoration site. For example, the “bluestems” can include native
grasses such as seacoast bluestem or big bluestem, but also
include non-native grasses such as the more invasive cultivars
of Kleberg bluestem or King Ranch bluestem.
Key points to consider when purchasing seed:
• Did the seed come from a local wild population, or was it
commercially produced?
• What is the origin of the original parent seed?
• What is the pure live seed (PLS)?
• What is the purity of the seed (how much inert material,
non-natives, and weed seed are present)?
Collecting native seed with a hand-held seed stripper
63

Types of Seed
Types of seed are based upon verification of species and source, and approved Texas Department
of Agriculture guidelines.
Selected Texas Native Germplasm (green label): Seed, seedlings, or other propagating materials
from untested parentage of rigidly selected native plant material stands that have promise, but not
proof, or genetic superiority.
Source Identified Texas Native Germplasm (yellow label): Seed, seedlings, or other propagating
materials collected from natural stands, seed production areas, or seed fields where no selection or
testing of the parent population has been made.
Cultivar: A variation of a species that has been chosen through breeding or selection. The use of
cultivars is questionable in native plant restoration because they may not maintain the genetic and
biological diversity of the species as a whole; they should be used with discretion.
Kind: One or more related species or subspecies, which are labeled singly or collectively under
one name (example: oats, bundleflower).
Variety (cultivar): A subdivision of a kind characterized by growth, yield, plant, fruit, seed, or other
characteristics, by which it can be differentiated from other plants of the same kind.
Certified seed: Seed that has been inspected by the Texas Department of Agriculture and meets
approved guidelines for the species. It is sold with a blue tag that lists information such as the
variety name, germination, weed seed percentage, seed lot number, and source of production.
Site Adapted Custom Seed Collections: Seed collections that have been custom harvested
by seed companies; some companies offer native harvested mixes, and also harvest seed from
customer specified sites.
Custom Mixes: Combinations of seed from more than one plant species that have been mixed by
seed companies to meet the preferences of customers.
Seed Quality
Quality of the seed that you are purchasing is very important.
characteristics can be found on seed tags.
Details regarding key seed
PLS (pure live seed) %: Percentage of the product that is made up of viable seed where PLS =
(% Germination + % Dormant Seed) x % Purity.
Germination %: The amount of seed that will grow under favorable conditions.
Purity %: Quantity of the desired product that is present in the bag.
Inert matter: Dirt, chaff, sticks or other non-seed material within the bag.
Dormancy: The amount of seed that will require longer periods of time to germinate.
Weed seed: The amount of non-target species seed that is contained in the bag.
Native or Non-native (pending Texas Department of Agriculture final seed-tag regulations): Seed
from a species that is not native to the region, usually introduced from another country.
Origin or lot number: Indicates the seed source, and can be used to trace the specific
characteristics about its origin.
64

Collecting Wild Native Seed
Hand collecting native seed
Before You Start
Consult with seed
Once you determine which species to collect, it is time to locate sites
experts when deciding
that contain sufficient quantities of the species you are interested in
upon which seed type
collecting. Seeds should be collected close to the restoration site in
to use.
order to obtain a similar “ecotype.” These plants are adapted to a
particular local environment, and will often thrive better than plants from
other regions.
Collection Timing
• For locating plants for collection, it is easiest to identify species during the
flowering stage. You can mark located plants with stakes, flagging tape, or GPS
coordinates to relocate them after they have gone to seed. Scouting for and
marking plants, especially forbs, while they are flowering will make them easier
to find once they have gone to seed.
• Rainfall in South Texas is usually the most important factor influencing whether
seeds will have good embryos; try to pick seed in years, seasons, or areas of
sufficient rainfall.
• After plants flower, periodic surveys of the selected plant communities will help determine the
harvest time.
• Picking seeds too early will result in the collection of immature seeds that will have low seed
viability. Also, because some seeds fall from the seed-head quickly after maturity, most seeds
could be missed if picked too late.
65

ETHICS OF HARVESTING WILD SEED
Do not collect
seed on federal or state
property without permission,
and do not collect
endangered or
It is critical to obtain landowner or land manager permission if collecting
on property other than your own. In particular, most state or federal
agencies require special permits to collect seed.
While restoring or improving another site, ensure that the source
population is not damaged by over-harvesting seed. If a source site is
threatened plants! scheduled for clearing by development or cultivation activities, seed can
be opportunistically collected from the source before it is destroyed. On
other areas, however, it is best to avoid harvesting all, or even most, of the
seed for each species on the site because many native species do not set viable
seed every year, and may have low germination. When choosing a collection site, look for a wellestablished
plant community of your target species. It is best to consult a restoration ecology
professional, or thoroughly research the species you plan to collect to ensure the population will
not be adversely impacted.
Many native species have indeterminate inflorescences where the flower stalk continues to grow
with prolonged flowering; it may require several collecting trips to effectively collect seeds for these
species. It’s important to pick mature seeds from healthy plants to help ensure superior offspring
with similar qualities.
Seed Maturity
Hairy grama (Bouteloua hirsuta) Indian blanket (Gaillardia pulchella) Desert Yaupon (Schaefferia cuneifolia)
Grasses
Mature seeds are
typically dry and hard,
and have separated from
the rachis or stalk.
Seed maturity can
sometimes be determined by using
the “hard dough” test. Moisture in
immature seeds is higher than in ripe
seeds, so most mature seeds will not
easily be squashed with
your fingernail.
Broadleaf Herbaceous
Plants
Mature seeds are typically
dry and hard, and loosen
easily from the pods,
capsules, or flower heads.
Shrubs & Trees
Fleshy seeds should be fully
ripe, and should be easy to
remove from the stem.
66
Checking a coma (Bumelia
celastrina) seed for ripeness

Seed Quantity
Before going into the field, decide how much seed to collect. Some of the factors to consider are
• Size of the area you wish to plant;
• Number of container-grown plants you wish to grow;
• How much seed is available for collection;
• Available equipment and labor;
• Time available for collection;
• Availability of storage space.
Because wild harvested seed often has low seed viability, collect more seed than has been calculated
for the restoration plan. Collect seed from a number of different plants to ensure diversity.
If collecting seed for South Texas Natives or one of the USDA-NRCS Plant Materials Centers, it is
best to collect approximately 1/4 pound of seed, if possible, but smaller collections are accepted. If
possible, seeds should be collected from a minimum of 30–50 plants.
HELPFUL TIP
If harvesting mechanically for a mixed collection of seed, it’s easiest to select for the predominant species
desired for the mix, and harvest when the majority of the seed is ripe. Because mixed seed is difficult
to seperate and clean, the mix will have to be planted as a combined mix, or sent to a seed-cleaning
facility.
Mechanical Seed Collection
Hand-held seed stripper
Mechanical harvesting can be economical for large-scale
collection efforts of both single species and mixed collections.
Machines that are used to harvest prairie seed can be
categorized into 2 general groups: (1) small machines such
as portable seed strippers, and (2) large machines such as
combines.
Native Seed Strippers
Hand-held Seed Stripper: This machine utilizes a rotating reel
that combs or sweeps seed heads off stalks, and then collects
and contains seeds in a detachable “hopper.” This
type of equipment is lightweight, and has a minimal
impact on the prairie landscape because it leaves
the plant stalks intact for bird nesting cover and
prescribed burns. It is useful for collecting seeds
from selected patches of plants, in areas of limited
access, or on rough terrain. These machines are
relatively inexpensive, efficient, and simple to
operate.
Harvested seed mix
67

Pull-type Seed Stripper: This machine is
similar to the design and function of the handheld
seed stripper, but is pulled as an implement
behind a tractor or all-terrain vehicle. The seedstripping
mechanism consists of a nylon bristle
brush powered by a 5-horsepower gasoline
engine mounted on the unit. The brush sweeps
seed into the hopper leaving plant stalks intact and
erect. The machine’s adjustable height, brush speed,
and brush angle allows for the harvest of various prairie
types and species.
Pull-type seed stripper
Flail-Vac®: The Flail-vac is an implement that is mounted on the front of a tractor, and uses a
spinning brush to create a vacuum that pulls the seed head off the stalk. Similar to the handheld
seed stripper, seed is then deposited into a self-contained hopper. These machines can be
operated on somewhat rough terrain, and are relatively maneuverable.
Larger Machinery
Combines: Combines are feasible for collecting native seed on large sites that are easily accessible.
Most combines are designed to harvest heavy-seeded agricultural crops, but modifications to the
headers and cutting height can increase efficiency for more lightweight, native seed. Unfortunately,
combines are expensive to purchase and maintain; however, services can be easily contracted.
Hay Equipment: Common field mowers can be used to harvest short grasses because of their
ability to cut closer to the ground. A sickle bar is also a good option for most types of grasses
and forbs. Because the sickle lays the hay over to one side, raking the hay into a windrow is not
necessary.
Flail-Vac seed stripper
68

The type of baling equipment used will be dependent on
the spreading technique. If using manure spreaders
or blowers to spread the hay, then a square baler
will be most practical. If planting in strips, however,
a baler that produces large round bales will be more
feasible.
Hand Collection
Activities
such as windrowing and
hay-raking can cause seed loss due
to shattering; therefore, it is important to
minimize hay handling. While the hay will take
longer to cure, more seed will be salvaged. If
handling cannot be avoided, it should be done
at night when evening dew will help to
minimize seed shattering.
Most native species do not grow in pure stands. Because of South
Texas’ topography and diverse brushy vegetation, seeds may have to be
collected by hand or with other small collection devices. Here are some helpful
tips:
• Fruits should be kept in a ventilated plastic bag and kept cool in an ice chest until the
flesh can be removed from the seeds.
• Non-fleshy seeds should be collected and stored in small paper bags. Do not use plastic
or wax-coated paper; this can cause mold to form because of trapped moisture.
• Remove excess litter using hardware cloth, or a fine wire mesh mounted on a wooden
frame.
• Collected seed should be labeled with the collection date, collection location (including GPS
coordinates), common name of the species, the scientific name (genus and species), and
collector’s name. Any descriptive notes associated with the plant or site such as slope, soil type,
location of plant (shade vs. sun), and the association of other plants growing nearby can be
useful information for future reference.
You can also save time by collecting
seeds in animal droppings. Seeds
are often naturally “scarified” by
the animal’s digestive tract, and may
germinate more easily. Just be sure
to wear gloves!
Texas persimmon in
animal dropping
Collecting fall witchgrass seed, Webb Co., TX
69

70
Notes

Drummonds skullcap (Scutellaria drummondii) and Drummonds Phlox (Phlox drummondii)
SEED PROCESSING
AND STORAGE
71

SEED PROCESSING AND
STORAGE
Seed Processing
Proper cleaning and storage techniques can help maintain seed viability over
extended periods of time. In addition, mature, unblemished seeds will remain
viable longer than immature or damaged seeds. Storage conditions such as
temperature and moisture will also affect the longevity and viability of seed.
Cleaning Fleshy Fruits
Most fleshy fruits need to be cleaned immediately after arriving from the field to prevent
seeds from degrading or spoiling if storing seeds for later planting. If necessary, seeds can be
briefly stored in a refrigerator until they can be cleaned.
Some species have “mummified” seeds that can be planted with the skins intact.
The mummified fruit may contain inhibitors that may cause somewhat of a delay in
germination, but is meant to provide a nutrient-rich substrate for microorganisms
during this critical period. After washing, mummified fruits should be spread on trays
or screens and dried in the sun. Examples of seeds that can be planted without
cleaning are grasses, non-fleshy forbs, and some fleshy fruits if planted immediately
following harvest.
Small amounts of fleshy fruit can be hand-squeezed or mashed by a rolling pin, fruit press, or by
rubbing the fruit against a screen with a brush. If the fruit clings to the seed, allow some drying
time, then rub the seeds on a screen until the dry matter flakes off.
Small-seeded fleshy fruits can be macerated in a blender, but use short, low-speed intervals to
avoid damaging the seeds. Run water over the seeds in a series of different sized colanders;
adding a paper coffee filter into a second colander will help to catch seeds.
Hand Threshing and Scalping Brittle
Seeds
Some seeds have brittle inflorescences or coverings (such as
pods) that can be broken away by hand threshing. The collected
material can be placed in a cloth sack or inside a cut length of
inner tube, and crushed/rolled until the dry matter has broken off.
A coarse screen is also a good tool, used with a gloved hand, plastic
brush, or rubber paddle.
Be familiar with the
germination requirements of
the seeds you are cleaning
- different processes could
affect their viability!
After threshing, clean the remaining litter by using 2 different sizes of screen. The first
screen allows the seeds to fall through and stop the coarse trash; the second will stop
seeds and fine trash from passing.
73

Mechanical Threshing and Cleaning
For larger seed-processing projects, hand threshing may not be adequate. Although somewhat
costly, machines such as the hammer mill, seed scalpers, seed screeners, de-bearders, seed
blowers, and seed separators can be very effective.
Contact a USDA-NRCS Plant Materials Center or commercial seed growers for more information.
Drying Seed
Drying techniques and moisture levels can vary and
depend on the type of seed in question. For example,
moisture is not as harmful to hard-coated seeds (such as
many legumes) because they are impermeable to water.
While most seeds are “desiccant-tolerant” and can be
dried, some seeds are “desiccant-intolerant,” and cannot
be dried; these include seeds with large cotyledons, oaks
and buckeyes, and most aquatic plants. “Recalcitrant”
seeds (another type of desiccant-intolerant seed) cannot
survive drying and freezing; these include certain tropical
fruits, palms, and most citrus. It is important, therefore, to
be familiar with the germination requirements of collected
seeds, and adapt drying and storage requirements
accordingly.
Methods
Because high humidity is not conducive for air-drying in
South Texas, seeds can be slowly dried in an oven at
Using a blower to clean seed.
approximately 100° F. To prevent seeds from drying too
fast, dry the seeds for 1-2 days at approximately 65° F, and then gradually
increase to 100° F until dry.
Seeds can be air-dried if humidity can be controlled between 20-40%
using a dehumidifier. A light bulb or low-level heat lamp can also be
used, but take care not to allow seeds to dry too fast.
Seeds can be placed in a small cloth bag, and sealed in a jar that
contains an equal amount of silica gel or fresh powdered milk; allow
8-12 days for small seeds, and 12-16 days for larger seeds.
These fruits can be stored for up to 6 months in cold storage (between
34-41° F). Desiccant-intolerant seeds must be kept moist, and can
only be stored for short periods (less than 3 months). After cleaning,
layer seeds on some moistened sphagnum moss in a non-airtight
container; seeds will require air for continued respiration. These fruits
can be stored for 2-3 months in cold storage (between 34-41° F), but
afterwards viability will begin to decline.
74
Drying seed using silica

Most fleshy seeds (berries, drupes, and pomes) will enter a stage of dormancy after being fully
dried. To keep this from happening, layer seeds in a damp, airtight container (after cleaning) along
with some moistened sphagnum moss.
Grass and forb seeds harvested at full maturity should require little to no drying time. When seeds
are unripe and green, some air-drying will be required, and can be done on screens or tarps. The
seed should be safely stored in a cool (65-75° F), dry facility in porous sacks (paper or cloth).
Storage
Long-Term Seed Storage
Moisture and temperature are the 2 most important factors affecting seed longevity. For the most
part, seeds deteriorate with high moisture; for every 1% reduction in seed moisture, a seed’s
storage life is doubled. If the seed’s moisture content is above 30%,
non-dormant seeds will germinate. Seeds also deteriorate with high
temperature; for each reduction of 10° F, a seed’s life is doubled.
Ideally, seeds should be stored in a climate-controlled facility
such as a seed vault after being properly dried. These facilities,
however, are expensive and not readily accessible to the general
public. The following are some practical methods for long-term seed
storage (excluding desiccant-intolerant seeds):
Dormancy
is a mechanism that
helps plants postpone
germination until
environmental conditions
are favorable
You can reactivate
silica gel in your oven at 250 o F until the
beads turn deep blue. Do not place seeds
in direct contact with silica; it may damage
seeds. Place containers in a secure freezer
(i.e., below 32 o F) where seeds can endure
long-term storage.
• Dried and cleaned seeds should
immediately be placed in moistureproof
containers; do not allow seeds to
regain moisture because the formation
of ice crystals after freezing will cause
damage.
• Place silica gel in a porous cheesecloth or nylon
stocking in the bottom of each storage container to
absorb moisture. Use 2 lbs of dry silica gel to 10 lbs
seed. The indicator silica gel will change from blue
to pink when the silica needs to be replaced.
A good rule of thumb for refrigerated storage is that the combination of temperature
and relative humidity should not exceed 100° F. So, for example, if the temperature
is 50° F, then the relative humidity must remain below 50%; if the temperature is
60° F, then the relative humidity must be maintained at 40% or lower.
HELPFUL TIPS
• Use only clean containers for newly harvested seed.
• To minimize insect damage, Diatomaceous Earth (DE) can be used to cover the surfaces of the seeds;
simply sprinkle the DE over the seeds, and mix gently; be sure to wear gloves because DE is abrasive
to skin.
• In worst-case scenarios, seeds can be fumigated; this is somewhat impractical due to the toxicity and
hazards of pesticides and applicator’s license requirements.
75

Protection from Insects
The best way to control most insects is by storing seeds at the proper moisture and temperature
levels. Freezing, In those plant species unaffected by low temperatures, also restricts pest
activity.
Native Hay Storage
Seed hay should be stored away from exposure to the elements, and protected from rodents. In
addition, make sure that hay is dry before baling; moist hay may heat up enough to kill some or all
of the seed. The hay seed can remain viable for several years if stored properly. Most native seeds
have some type of dormancy, so holding hay for a certain time could increase germination of some
species. Little information regarding native seed hay storage is available at this time, so exact time
frames and specific requirements are speculative.
Bulk or Bagged Native
Seed Storage
Processed seed material should be
stored in a barn or shed where it
can be protected from rodents and
the weather. Similar to seed hay,
storage requirements are unknown
for native wild harvested seed.
Check with your seed dealer to
determine storage requirements for
commercial seed. It is best not to
take delivery of the seed you have
purchased until you are ready to
plant in order to avoid unnecessary
storage expense and time.
Seed storage vault, E. Kika Garza Plant Materials Center
USEFUL REFERENCES
Nokes, J. 2001. How to Grow Native Plants of Texas and the Southwest. University of Texas Press, Austin, Texas.
Young, J. A., and C. G. Young. 1986. Collecting, Processing and Germinating Seeds of Wildland Plants. Timber Press,
Inc., Portland, Oregon.
Florabank – http://www.florabank.org.au/
Reforestry, Nurseries, and Genetics Resources – http://www.rngr.net/Publications
Department of Forestry, Auburn University – http://www.forestry.auburn.edu/sfnmc/class/handbook/ch5.html
76

Wildflowers, Medina Co., Texas.
77

78
Notes

Wooly globe mallow (Sphaeralcea lindheimeri)
SEED APPLICATION
79

SEED APPLICATION
Seed application methods vary with the type of seed (wild harvested seed or hay, or commercially
purchased seed), and with the type of terrain at the restoration site. Once the seedbed is prepared,
seed can then be drilled or spread using a variety of implements and techniques.
SEEDING RATES
If planting commercial seed, use the PLS value printed on the seed tag. If using a wild native harvested
mix, use the estimated PLS of the dominant species in the mix.
To calibrate the seeder, calculate the amount of “bulk seed” based on the PLS value (i.e., the total amount
of viable seed). Divide total bulk pounds by total acres, giving the planting rate/acre.
Bulk seed (lbs/ac) = Desired rate of seed in PLS (lbs/ac)
% PLS of current seed lot
Broadcast Seeding
Broadcast seeding is used to scatter seeds across
large areas by air or ground. In ground application
methods, a seeder box or spreader is attached to a
tractor, pick-up truck, or ATV, and can be operated
by one person. Seeds can also be dispersed by
airplane or helicopter. Seeds are evenly dispersed
through a spreader attached to the bottom of a
hopper in the aircraft. In general, broadcast seeding
is less expensive than other complex, mechanized
techniques, and requires less time and effort. In
addition, broadcast seeding can be more effective
than other methods during times of low rainfall;
seeds can be quickly dispersed to take advantage of
unexpected or isolated showers. Lower germination
rates usually result because some seeds are left
uncovered and exposed to the elements. Broadcast
seeding should not be used on seedbeds that are
rough and cloddy or crusted over. Broadcast seeding
is commonly used in South Texas to restore areas
with rough or brushy terrain, or when fields are too
wet for cultivation. If possible, rollers, packers, or
drags should be used on broadcast seeds to ensure
good contact with the soil, and to lightly cover the
seed.
Seed broadcaster
81

Drill Seeding
Consult with other
consumers and ask
Seed drills are machines that sow seeds in well-spaced for references when
rows at specific depths. Rangeland seed drills are based puchasing seeding
on conventional drills, but have specific modifications for equipment.
dealing with the planting requirements associated with planting
small, fluffy, and chaffy native seeds. Some modifications may
include agitators housed in seed boxes, special tubes for ease of seed
flow, and various planting and covering modifications. The seeds of many native
grasses are covered in small hairs, or have awns that easily entwine and adhere to
one another. Seed box agitators keep seed from becoming compacted or attached to
one another. These agitators come in a variety of forms, and keep seed flowing into the seed tubes
at a constant rate.
Some rangeland seed drills are equipped with special
tubes that help seeds to travel from seed boxes to
planting discs more easily. These tubes are pliable for
drill movement and flex over rough rangeland terrain;
they are easy to disassemble for obstruction removal.
Planting and covering modifications on rangeland
seed drills are adjustable to allow for shallow planting
depth requirements, and for planting on uneven
terrain. Many rangeland drills are designed to plant
with no-till capabilities, which can be helpful in areas
where disturbance might cause excessive erosion or
weed invasion. Manipulating the planting depth also
allows better control over germination. Generally,
drill seeding requires less seed and produces higher
germination rates than with broadcast seeding.
Discretion and research should be used when
considering the purchase of rangeland seed drills.
Many have reliable modifications that perform well.
On the other hand, some drills have undependable
modifications that require further work and add
increased costs of frustration and labor to an already
significant financial investment.
“No-till” rangeland drill
TIPS FOR BROADCAST SEEDING
• Always broadcast cross-wind.
• Calibrate equipment based on rate of output and speed of travel.
• Check manufacturer’s instructions for your equipment.
• Plant a test strip or plot to determine if the seeding rate is accurate and sufficient, and make necessary
adjustments to achieve desired planting rate.
• It may be necessary to add carriers such as rice hulls or crimped oats to help with seed flowability.
82

Pitters, dikers, imprinters, aerators, and other similar devices are used on rangelands or arid regions
where preparing a seedbed is impractical. These implements create small depressions (basins) in
the soil that help capture moisture in low rainfall areas, and also help with soil aeration and water
infiltration. Seeds are then broadcast either by ground or aerially following the treatment. Some
imprinters can be fitted with a seed bin that presses seeds into the walls of the depressions while
imprinting. Rainfall then erodes the soil into the basin to cover the seed. These treatments seem to
work best in loamy textured soils that are loose enough to form a basin, but firm enough to maintain
the basins for some time after seeding.
Seeding Cultivated Land
Before broadcast or drill seeding in cultivated fields, coarsely textured soil must be dragged or rolled
to lightly pack the soil before planting. Some seeders are designed to simultaneously perform
this task, and gently press seeds into the soil with rollers as seeds are dropped from the seed
opening.
For broadcast seeding, packing is generally only necessary to press, but not bury the seed into the
soil for good soil-to-seed contact. Most native seed is small so do not bury the seed. Rubber tire
rollers designed for agricultural use are very effective in firming the soil and enhancing soil-to-seed
contact.
With fine-textured soils, packing should not be conducted before planting. This procedure hardens
soil, leaving broadcast seeds on the soil surface and exposing them to dry air and birds.
SEED FLOWABILITY
Seed enhancements, such as seed coatings, can improve flowability through planting equipment by
altering the shape of the seeds. For example, fluffy or chaffy seeds can be altered into small pellets,
which flow more smoothly through seeders. Additionally, seed can be de-awned or de-bearded with
specialized seed cleaning equipment which removes the appendages by abrasion.
There are a variety of coatings, including water absorbing polymers, that may enhance germination
success, or microbial inocula that are critical for germination of some plants. Other coatings can provide
protection from stress conditions, rodents, and birds, and also help increase nutrient availability. In some
cases, coatings can provide protection from seedling diseases, and the negative effects of fertilizers and
herbicides.
With respect to South Texas native plants, most seed coating techniques are experimental and warrant
further study. Contact a regional USDA-NRCS Plant Materials Center or seed dealer to ask about any
advances or pertinent information regarding the species with which you are working.
Chaffy seed before
coating
Coated seed
Silver bluestem (Bothriochloa laguroides)
83

Thin paspalum (Paspalum setaceum) False rhodesgrass (Chloris crinita) Indian blanket (Gaillardia pulchella)
A challenge in native plant restoration is the diversity of seed types. Native plant seeds vary in size, shape,
and texture; their planting ease, or “flowability,” may not be conducive for the conventional seeding methods
used in today’s agronomic industry.
Seed Drill Calibration
Seed drills require calibration to ensure an accurate seeding rate. Factors to consider when
calibrating include seed size and weight, mixture, moisture content, degree of processing, tire
slippage, and amount of trash present.
SHOP DRILL CALIBRATION
Determine the desired seeding rate using the current lot of seed
Determine the pounds per acre of bulk seed that is needed using the current seed lot.
Bulk seed of current lot needed (lbs/ac)= Desired rate of seed in PLS (lbs/ac)
% PLS of current seed lot
Set Drill
Set the drill to the seeding rate desired.
Collect Actual Seeding Rate Of Drill
Jack the driving wheel of the drill up, turn 20 revolutions, catching the seed from one run of the seed
drill in a bag or sack. Weigh sack to determine pounds of seed collected.
Determine Wheel Circumference
Measure the circumference around the outside of the tire with a measuring tape (in feet)
Determine Strip Length
Strip length (in shop) = 1.1 (No. of revolutions X wheel circumference (ft))
NOTE: The 1.1 is to compensate for wheel slippage in the field.
Determine Current Seeding Rate
Using the answers for the above steps, calculate the following equation:
Seeding rate in bulk seed (lbs/ac) = 43560 ft²/ac X lbs seed collected
Drill width (ft) X Strip Length (ft)
If both sides of the equation are equal, the drill is calibrated. If unequal, the drill needs to be
adjusted, and the process repeated until the drill is calibrated.
84

SHOP DRILL CALIBRATION EXAMPLE
A seed mixture needs to be planted at a rate of 15 lbs PLS per acre. Your current seed lot has a PLS of
82%. After jacking up the driving wheel and turning for 20 revolutions, 1.2 lbs of seed were collected. The
drill is 15 feet wide and the tire circumference is 6 ft.
Step 1 - Determine the desired seeding rate using the current lot of seed:
Bulk seed of current lot needed (lbs/ac) = Desired rate of seed in PLS (lbs/ac)
% PLS of current seed lot
Bulk seed of current lot needed (lbs/ac) = 15 lbs/ac PLS needed
82% PLS
Bulk seed of current lot needed (lbs/ac) = 18.3 lbs/ac
Steps 2 through 4: Completed
Step 5 - Determine strip length:
Strip length (in shop) = 1.1(No. of revolutions X wheel circumference (ft) )
Strip length (in shop) = 1.1(20 revolutions X 6 ft)
Strip length (in shop) = 1.1(120 ft)
Strip length (in shop) = 132 ft.
Step 6 Determine Current Seeding Rate:
Seeding rate (lbs/ac) = 43560 X lbs seed collected
Drill width (ft) X Strip Length (ft)
18.3 lbs/ac = 43560 ft²/ac X 1.2 lbs seed collected
(15 ft X 132 ft)
18.3 lbs/ac = 52272 lbs
1980
18.3 lbs/ac = 26.4 lbs/ac
Because the equation was not equal, and the drill was putting out 8.1 lbs/ac too much, the drill needs to
be set lower and recalibrated until the equation is equal.
FIELD DRILL CALIBRATION
Determine the Desired Seeding Rate Using the Current Lot of Seed
Using the current seed lot, determine the pounds per acre of bulk seed that is needed.
Bulk seed of current lot needed (lbs/ac) = Desired rate of seed in PLS (lbs/ac)
% PLS of current seed lot
Set Drill
Set the drill to the seeding rate desired.
85

Collect Actual Seeding Rate of Drill
Travel a distance of 100 ft with tractor and drill, collecting seed from one run. Weigh the amount
of seed collected.
Determine Strip Length
Strip length (in field) = The distance driven to collect seed, this is usually 100 ft
Determine Current Seeding Rate
Seeding rate (lbs/ac) = 43560 ft²/ac X lbs seed collected
Drill width (ft) X Strip Length (ft)
Plug the numbers already determined into the equation. If both sides of the equation are equal, the
drill is calibrated. If the numbers on either side of the equation are not equal, the drill needs to be
adjusted and the process repeated until the drill is calibrated.
FIELD DRILL CALIBRATION EXAMPLE
A seed mixture needs to be planted at a rate of 15 lbs PLS per acre. Your current seed lot has a PLS of
82%. A tractor with a 15 ft drill was driven 100 feet while collecting seed; 0.4 lbs of seed were collected.
Step 1 – Determine the desired seeding rate using the current lot of seed:
Bulk seed of current lot needed (lbs/ac) = Desired rate of seed in PLS (lbs/ac)
% PLS of current seed lot
Bulk seed of current lot needed (lbs/ac) = 15 lbs/ac PLS needed
82% PLS
Bulk seed of current lot needed (lbs/ac) = 18.3 lbs/ac
Steps 2 and 3: Completed
Step 4 – Determine strip length:
Strip length (in field) = The distance driven to collect seed
Strip length = 100 feet
Step 5 – Determine Current Seeding Rate:
Seeding rate (lbs/ac) = 43560 ft²/ac X lbs seed collected
Drill width (ft) X Strip Length (ft)
18.3 lbs/ac = 43560 ft²/ac X 0.4 lbs seed collected
(15 ft X 100 ft)
18.3 lbs/ac = 17,424 lbs/ac
1,500
18.3 lbs/ac = 11.6 lbs/ac
Because the equation was not equal, and the drill was putting out 6.7 lbs/ac less than required, the drill
needs to be set at a higher rate and recalibrated until the equation is equal.
86

Hydroseeding and Mulching
Hydroseeding and mulching are alternatives for rapid establishment of vegetation in habitat
restoration or erosion control efforts. While hydroseeding can sometimes cost more than other
techniques, there is little impact from heavy machinery to the seedbed; this provides a more stable
system for establishing plants.
Hydroseeding is a technique where water, seed, and fertilizer are
sprayed over an area. Hydromulching is similar to hydroseeding
except that mulch material and tackifier are included in the spray
solution. Mulch material may consist of a paper, wood fiber, or bonded
fiber matrix.
Be careful not
to bury seed
too deeply!
Mulches protect the soil from erosion, and help to regulate soil temperature; they
also help retain water and protect the soil from raindrop splash. Each type of mulch
has different capabilities. Paper and wood fiber mulches perform similarly on flatter
surfaces, but require lighter rates or thicknesses. When higher rates are used, paper
mulches can form a barrier that, in some cases, can hinder seedling emergence.
Wood fiber products generally do not produce this problem. Bonded fiber matrix products
perform similarly to both products on flatter surfaces, but are best used on steeper slopes where
erosion is a concern.
Hydroseeding Rates
Paper and wood fiber hydroseeding rates differ depending on the site and soil types. They are as
follows:
• Sandy soils with < 3:1 slope requires 2,500 lbs/ac of mulch.
• Sandy soils with > 3:1 slope requires at least 3,000 lbs/ac; in steep sandy soils, a bonded fiber
matrix or a soil retention blanket should also be used.
• Flatter clay soils of < 3:1 slope requires 2,000 lb/ac.
• Steeper clay soils of > 3:1 slope requires 2,300 lb/ac.
• Bonded fiber matrix materials can be applied at rates of 3,000 to 4,000 lb/ac. At higher rates,
some of the bonded fiber matrix materials can be used in lieu of a soil retention blanket.
Tackifiers
One of the most important factors when performing a seeding operation is to prevent movement
of the seed and soil, and this can be accomplished using a tackifier. Basically, a tackifier is a
type of glue that holds mulch, seed, and soil together. There are a variety of tackifiers on the
market including latex binders and guar products. All work well, but if working with a state or
federal agency, most must be approved before use, and should be applied using manufacturer’s
recommendations.
Equipment
Hydroseeding equipment consists of a specialized tank, hose, pump, agitator and hand applicator,
and is set up for spraying light mulch rates, seed, and fertilizer. The agitator (mechanical or jet)
keeps the mulch, seed, and fertilizer mixed properly for even distribution.
87

Hydromulching requires a higher
volume pump and stronger
mechanical agitator to keep the
thick slurry of mulch material
mixed. Higher pump capacity
is also needed to spray the thick
slurry through a longer hose to
reach inaccessible areas.
Water Requirements
Access to water is very
important when hydromulching.
Approximately 6,000 gallons of
water are required to seed one
acre. A nearby water source is
essential to minimize costs and to
complete a large task within a reasonable time frame.
Hydroseeding along a right-of-way
Application Techniques
To prepare a site for hydroseeding, the upper 6” soil layer should be tilled, and all large clods broken
up. Irrigation of the seedbed before and after the seeding operation is also recommended. Seed
and fertilizer can be applied simultaneously during a hydroseeding operation, but even distribution
of the material is important. Adding a small amount of dye will give the slurry a green color,
and will help with even distribution because of the green demarcation of the applied area. Seed
should be added no more than 30 minutes beforehand to the spray solution just before application;
seed allowed to soak in water too long may cause premature germination and, if applied to a dry
seedbed, could result in seedling loss.
One method for hydromulching involves a 2-step process, where seed and fertilizer are applied
before the mulch application. This ensures a high rate of seed/soil contact, keeping the mulch
from tying up the seed. Mulch is then carefully applied to avoid disrupting the soil surface, thereby
preventing soil erosion problems. Mulch should be allowed to gently fall on top of the soil, creating
a continuous mulch surface. Another method is to simply apply all of the materials at once. This
method is somewhat faster, but seed/soil contact may be somewhat diminished.
Wild Harvested Native Hay Application
Wild harvested native hay is spread either manually or with hay mulching/blowing equipment (such
as a manure spreader or hay blower), followed by light tillage used to cover the seed and anchor the
hay. A hay buster is the most effective method for distributing native seed hay if the terrain permits.
When using a manure spreader, it is most practical to have the hay stacked on a trailer that follows
and continuously feeds the hay into the spreader. A hay mulcher/blower is very effective, but might
not be readily available in most areas. This machinery can also be expensive, and may require
contract services to perform the application. Hay blowers are particularly effective for applying hay
to slopes and rough terrain. Hay can also be spread by hand where labor is available.
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Spreading rates depend on factors such as the amount of hay and seed harvested, the volume
of the species harvested, the size of the area to be planted, and the method of application. As a
general rule, it is difficult to spread less than 500 lbs of material/acre with a manure spreader. When
planting hay harvested from short grasses such as red grama, buffalograss, or curlymesquite, it
may be necessary to augment the amount of bulk material with some other type of clean straw to
facilitate the application. In any case, it is better to over-spread than to under-spread to increase
the chances of seedling establishment.
Hay should be anchored, or pushed into the ground. It may be necessary to apply hay in sections
and anchor it immediately to keep it from blowing off the application site; this will help to cover the
seed and hay at varying depths. Aerators, imprinters, packers, disks, or harrows are effective tools
for covering and anchoring the hay, but livestock trampling can also be used to create the same
effect. If using a disk or harrow, the implement should be set to run level and very shallow and
with the blades straight, not angled. Because most native seeds require little soil cover, coverage
depths should vary from the surface to less than an inch deep.
Anchor the hay, taking
care not to bury it too
deeply.
LITERATURE CITED
Pratt, M., and G. A. Rasmussen. Drill Calibration, Utah State University Extension,
http://extension.usu.edu/files/natrpubs/range2.pdf.
Chapman, S. R., and L. P. Carter. 1996. Crop Production: Principles and Practices. W.H.
Freeman and Company. San Francisco, Calif.
Native seed hay
89

Notes
90
Spring thunderstorm, Jim Hogg Co., TX

Sideoats grama (Bouteloua curtipendula)
PROPAGATING AND
TRANSPLANTING
91

PROPAGATING AND
TRANSPLANTING
Why Propagate Native Plants?
Most native seed, especially those for shrubs and trees, are not readily available on the commercial
market. Because most seeds must be collected by hand or by crude mechanical means, prices
are high and availability is low. Growing your own transplants, purchasing transplants from a
nurseryman, or contracting an experienced commercial grower requires less seed than direct
sowing. Seedling establishment risks are also lower because transplanting
can be optimized to avoid the South Texas heat. Transplants provide
uniformity, and often grow faster than seedlings that were directly sown
as seeds. Furthermore, propagated transplants compete better than
newly germinated seedlings against aggressive weeds because they
have already developed root systems. For restoring vegetation on very harsh
environments such as saline sites, transplanting can be very effective because
salt levels may be too high for germinating seeds. Transplanted seedlings are
more likely to tolerate existing salt conditions because of their well-developed
roots. A drawback to transplanting is the potential high cost of labor and material
to grow and transfer the seedlings into the field.
Several processes are necessary to increase the chances of
native seedling survival and successful re-establishment at
restoration sites:
• Plan Ahead: Planning efforts should focus on seed collection, finding
suitable space, and a safe environment for initial propagation, site
preparation, timing of planting (weather, especially rainfall patterns),
planting techniques, and seedling maintenance.
• Water: One key factor to successful transplanting is adequate water
availability and conservation of soil moisture for newly transplanted
seedlings. Planning and installing field irrigation needs to be done well
in advance of transplanting.
• Be Familiar with the Species Requirements: It is
important to understand the specific environment in
which native species naturally grow. Restoration sites
must be as similar as possible to a species’ native habitat.
Duplicating this environment will involve knowing the
light (sun vs. shade), soil type, soil moisture, and soil
pH requirements of the species you are transplanting
before you start.
Awnless bush sunflower in 5”x1”x1” plant band
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Germination
Germination is the beginning of growth in a seed.
Seeds remain in an inactive, non-growing state,
until conditions are right for germination. All
seeds need water, oxygen, and proper temperature
in order to germinate; some seeds have special light
requirements as well.
Heartleaf hibiscus (Hibiscus cardiophyllus)
If appropriate facilities are available, seeds can be germinated
on the premises; this can be somewhat labor intensive
and costly, however. There are reputable companies
that provide efficient and cost-effective services of
germinating and preparing seedlings for transplanting.
Most companies use planting trays or receptacles
that will fit specific transplanting equipment, so
make sure the trays will be compatible to the type of
transplanter you will be using in the field. Techniques
and receptacles used for transplanting grasses, forbs,
and shrubs vary, and depend on transplanting methods, cost,
or time and labor constraints.
In order to determine how many seeds to plant, you will need to know the pure
live seed (PLS) of each species being used. Most commercial seed has the
germination rate information printed on the label, but if planting wild harvested
native seed, germination tests can be conducted through a seed-testing lab.
Most non-cultivated seed generally has a low germination percentage. To ensure
the highest germination count possible, seed should be
collected during peak maturity, and properly stored.
Seeds can be density graded to improve quality for
growing transplants. This process sorts out heavier
seeds, and removes lightweight, weak, and immature
seeds that may have a low germination potential.
Find out about and apply the specific germination and
planting requirements for each of the species that you
are transplanting.
Different species require different germination periods
and growth times to reach appropriate transplanting
sizes. While many seeds can be easily germinated
directly in the soil, some seeds may require specific
preparatory treatments to initiate the germination
process. By becoming familiar with the requirements of
each species or seed type, you will be able to incorporate
the timing and special equipment needs into your plan.
You can find germination
information on seed tags, at USDA-
NRCS Plant Materials Centers, on the
USDA website http://plants.usda.gov,
on the Native Plant Network website at
www.nativeplants.for.uidaho.edu/, or in
Appendix F of this manual.
Transplanting grass seedlings by hand
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Use 5 to 10 times more seed than you intend to transplant, because germination of some native
plant species can be difficult and some seeds have special germination requirements. This will
provide some room for error caused by infertile seeds, insect damage, and seedling mortality during
planting and transplanting.
Testing for Germination Rates
Germination experiments are used to determine the active germination of a batch of seeds under
optimal conditions. Accurate estimates are very important to the overall success of restoration
projects because seed germination is a critical factor in calculating seeding rates and PLS. Seed
germination is also an important characteristic to consider when selecting the amounts and types
of species to use.
The germination experiments should be
representative of the entire batch of seed,
so random samples should be taken from
the overall sample. Multiple repetitions from
the entire batch are also recommended to
ensure that the results are dependable.
Germination tests can be conducted through
commercial seed testing laboratories, and
can cost anywhere from $25-$75 per seed
batch. Germination chambers are used
to mimic natural light/dark and heating/
cooling cycles, and are commonly used in
a variety of research facilities. Equipment
types and sizes vary, and prices can range
from hundreds to thousands of dollars.
Checking on germination rates of seedlings
Field or home tests can also be conducted fairly easily using the following steps:
• Count out a known number of seeds from the entire batch of seed; 3 repetitions of 50
seeds/batch of seed are sufficient.
• Place seeds into waterproof, airtight containers that allow light exposure to the seeds. Place a
paper towel or water absorbent material in the bottom of the container, and saturate with purified
water (seeds should not float).
• Place containers and seeds into a sunny area. It is important to note that internal temperatures
of airtight containers can be up to 20° F higher than air temperatures. Monitor temperatures
and if temperatures exceed 85° F, move containers to cooler location. Under artificial lighting,
temperatures for standard germination tests generally range from 65–85° F, but check to see if
specific species temperature guidelines are available from seed labs or website publications.
Light/dark cycles should mimic the natural germination conditions for the species that you are
propagating. Standard light/dark cycles are generally 12:12 hours dark:light. As with temperature
requirements, check with seed labs or website publications for additional information.
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• Count the number of seeds that germinate on a regular basis throughout a 30-day period. While
checking seeds, add water as necessary to prevent seeds from drying out. Calculate the average
for the number of seeds germinated across repetitions; this will be the active germination value
for the seed batch. If results are highly variable, request a standard germination test through a
commercial seed testing laboratory.
Seed Scarification
Some South Texas native species exhibit “hard
seed” characteristics, and often require a breaking,
scratching, or softening of the seed coat. Naturally,
this might occur over time in the soil, or for some
species, in the digestive tract of an animal or with
fire. Blackbrush, honey mesquite, huisache, and
Texas ebony seeds, for example, will not readily
absorb moisture when planted under normal soil
conditions. Their seed coats are impermeable and
will not germinate until the seed coat is disrupted
or deteriorates naturally in the soil. Natural seed
scarification can be mimicked using mechanical or
chemical techniques that alter the seed coat and
allow germination to occur.
Scarifying a blackbrush acacia (Acacia rigidula) seed.
Most hard-seeded native legume seeds are easily scarified using techniques that can be
implemented with limited equipment and facilities. With scarification or germination techniques,
seeds should not be treated or soaked until a few days before planting; generally, seeds that have
been soaked should not be placed back into storage.
Aerated Water Pre-Treatment
Most seeds will benefit from an aerated water treatment, and few will be harmed by it. Soaking
is a good method for “priming” seeds for germination, and can be practical for a wide range of
restoration projects. Soaking seed in pure aerated water for 24 hours will cause germination in
significantly less time than with un-soaked seed. This technique helps seed absorb water, signaling
that the environment is adequately supplied with water for survival, thus triggering germination.
Aeration oxygenates the water and prevents seeds from drowning. Seeds can be soaked without
aeration for a few hours without causing harm, but aeration should be used for longer periods when
soaking many seeds at once. For aerated water pre-treatment, follow these instructions:
• Cut out part of the top of a clean 1-gallon water or milk jug with the handle attached, and fill 1/3
with tap water.
• Connect a piece of plastic tubing long enough to reach from an inexpensive fish tank aerator to
the bottom of the jug. Run the tubing through the jug handle, and anchor the end of the tubing in
the bottom of the jug using a fishing weight or a heavy washer.
• Pour the seeds into the jug, using no more seeds than can be covered by the water. Be careful
to place the jug and aerator on a counter, table, or the floor where the aerator will not vibrate off
the surface.
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• Leave the aerator running between 12–24 hours for most seeds, and up to 3 days for very hard
seeds such as those from ebony or blackbrush. Check the jug periodically to be sure water does
not foam up over the rim. Because some seeds produce abundant foam, placing the jug in a tray
will reduce clean-up time.
• Change the water at least once every 24 hours, and more often if it becomes excessively
discolored.
Hand or Mechanical Scarification
Hand scarification is practical with large-seeded legumes such as Texas ebony or mountain laurel,
but impractical for large quantities or smaller-seeded species. Seed coats can be gently cracked
with a hammer, filed with a metal file, or nicked with a knife to break open or weaken the seed coat.
Seeds can also be lightly scratched using sandpaper. For example, seeds can be placed in one
layer across a piece of coarse sandpaper, and rubbed with a sanding block or another piece of
sandpaper. Another simple method is to line a jar with sandpaper and vigorously shake seeds. It
is important to stop when the seed is thoroughly scratched, but before the seed coat breaks. For
large quantities, a bench-grinder can be used to scratch seeds by gently passing the grinder across
the surface of the seeds within a confined container. A gem polisher or rock tumbler is an effective
tool to scarify seeds. For larger batches, commercial electric seed scarifiers that move seeds
around a rotating drum with propellers or air are also available for purchase.
Boiling Water Treatment
The boiling water treatment is practical for moderate quantities of seeds, and is relatively safe. This
technique should be used for hard seed only. Also, be careful because seeds can be killed if boiled
for too long. The boiling water treatment will help the seed coat expand and contract, cracking it
slightly and enabling the seed to imbibe water. Follow these instructions:
• Fill a bowl with tap water and set it aside on a surface convenient to the stovetop.
• Place clean seeds in a strainer with handles that can be held while
submerged in boiling water.
• Fill a large pot with tap water to within 1” of the top; make
sure strainer can be held in the pot in such a way that the
seeds are covered by the water but are not able to float over
the rim of the strainer. A strainer which is only a little smaller
in diameter than the pot is best.
• Bring the water in the pot to a rolling boil.
• Dip the strainer with seeds into the boiling water and hold there for 15
seconds.
• Immediately remove and dip the strainer into cool water in the bowl and hold there for
15 seconds.
• Immediately remove and dip the strainer back into the boiling water for 15 more
seconds.
• Repeat this step until the seeds have been dipped into both the boiling and cool water 3 times.
• Seeds may be planted as soon as they are cool enough to handle.
A successful aerated
water pre-treatment will result in
“plump” seeds whose coats have
absorbed the water. If no absorption
takes place, then seeds should be
scarified using one of the more
intensive techniques listed here.
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Chemical Treatments
Chemical treatments can be very effective for seed scarification, but some restrictions apply. Acid
scarification requires caution because some acids can be highly corrosive, and are extremely
dangerous to eyes, skin, and clothes. Be sure to follow all safety standards, and to conduct your
work in appropriately equipped facilities. In addition, because seeds vary in permeability, treatment
time can vary from a few seconds to 6 hours. If left in acid too long, seeds can be damaged. Testing
is the best way to determine duration of treatment, and can be done by removing sub-samples of
seeds at set intervals and visually checking the thickness of the seed coats. When coats become
paper thin, treatment should be terminated.
Testing for Scarification Treatment Effectiveness
Test the treatment effectiveness by soaking the treated seed in pure aerated water for 24 hours.
Each seed should swell if the treatment was effective. If not, then it may be necessary to increase
the acid or boiling water treatment time.
Planting Containers
A variety of planting containers are available for different growth forms and planting strategies.
One of the most important objectives in propagating your seedlings is to develop a healthy root
system before transplanting into the field to ensure the highest probability of success. Below are
some guidelines for choosing the right container system for your propagation objectives.
Trays or Flats
Trays (or flats) are available in many different styles, and can be made of different materials.
Biodegradable trays or containers can be planted directly into the ground, so this eliminates the need
to remove transplants from containers. Plastic or polystyrene foam trays, however, are practical
because they can be washed and reused, and help to insulate transplants in colder temperatures.
Trays are also relatively inexpensive and save space; larger, more expensive containers can
sometimes be wasted because they use more potting medium, take up more table space, and not
all of the planted seeds will germinate. Each tray contains a number of cells that accommodate
individual seedlings as they germinate and grow. Because seedlings depend upon their roots for
obtaining nutrients from the soil, more roots can lead to a higher chance of establishment. It is
critical that the cell size be sufficient to allow root development.
HELPFUL TIP
Check to see if chemical treatments can be replaced using less hazardous techniques such as water aeration, boiling
water treatments, or mechanical treatments.
USEFUL REFERENCES
Websites with helpful germination and propagation information are:
• http://www.fs.fed.us/database/feis/plants/index.html
• http://plants.usda.gov/
Nokes (2001) provides a thorough reference for chemical scarification techniques that can be applied to many South
Texas native plant seeds.
98

HELPFUL TIP
Seed planting depth varies with species; check the literature or the nearest USDA-NRCS Plant Materials Center to find
the best recommendation. As a general “rule of thumb,” however, seeds should not be planted deeper than 2 times
their diameter.
Trays with smaller cell sizes may be less expensive, but may result in overall seedling losses for
deeper rooted perennials. Many trays are designed to fit into special holding racks and transplanters,
and are well-suited for medium to large scale transplanting operations of grasses and non-woody
forbs.
Before seeding, plant containers should be packed and moistened to allow the soil to settle; this
will also ensure good soil moisture throughout the container. After sprinkling the seeds into the
cells, more soil should be added and packed to cover the seeds with another ¼” layer to prevent
them from being washed out while watering. Between ½ - 1” should be allowed from the top of the
container as a reservoir for top watering. This level will vary based on the water absorption rate of
the potting medium; native soils will absorb water more slowly than commercial potting soils.
Plant Bands
Plant bands are perforated individual square tubes available in a variety of sizes, and can be planted
in the field by hand or with a modified tree planter drawn by a farm tractor. The design of the plant
band is conducive to good root establishment for woody forbs and shrubs, which generally have
longer and more extensive root systems than grasses. Individual containers
also provide better control of root aeration and moisture conditions for growing
seedlings. Most plant bands have no bottom or top, they conveniently
fit into plastic crates which facilitate handling and transport, and they are
biodegradable. However, labor costs for packing soil into plant bands is
significantly higher than for flats or trays. While the net cost of plant bands
may be more expensive than trays, seeding in plant bands may eliminate
the need to transplant seedlings (which can damage root systems) into
larger containers. Seedlings are transplanted directly into the ground
with the plant band still intact. However, because they deteriorate over
time and plants may outgrow the containers, their use is limited if plants
must be kept in greenhouses for longer than one year.
A healthy root
system in your
transplants is
critical!
Left to Right: 6 cell flat, 1”x1”x3” band, 1”x1”x”5” band, tube, and a 4”x4” container. There are numerous styles of containers
used to grow native plants.
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Greenhouse seedlings in flat trays
Crates can be used to hold a number of plant bands, which are useful for simultaneously packing
and maintenance. The soil mix must be firmly packed into the plant band by hand about ¾ full so it
will not fall through the open bottom. The soil mix is then troweled into the tops of the plant bands
in each full crate, and is shaken, tamped, or dropped to compact the soil. More soil mix is troweled
on top and shaken down, and repeated until the soil is about 1” from the top. After scattering the
seeds into the plant bands, cover with an additional ¼”, pack the soil again, leaving an approximate
¾ -1” from the top of the container. This will allow an adequate reservoir for top watering.
As the seedling grows and its roots emerge from the open bottom of the plant band, the roots are
air-pruned as they encounter the hard surface of the plant rack or nursery bench. This encourages
more root growth so the plant band eventually fills with roots. The square shape of the plant band
also encourages roots to grow down rather than around, and helps prevent root binding. When
the seedling and container are planted, the roots continue to grow out through the bottom, and the
cardboard gradually biodegrades in the soil. This reduces transplant shock and encourages deep
rather than shallow root growth. Furthermore, the enclosed area helps to retain moisture around
the roots of the transplant, helping it to become established. Special care should be taken to cover
the top of the plant band below the soil surface; if not, the cardboard can actually serve as an
evaporative wick and cause moisture loss to the seedling. If irrigation is available, the use of plant
bands may not be necessary.
100
Plastic Containers
Plastic potting containers are suitable for woody-stemmed forbs and shrubs, and are generally
used for smaller-scale projects where plants will be transplanted by hand. These containers can
be more expensive than trays or plant bands, and while they are somewhat impractical for larger
restoration efforts, they can be useful in smaller projects such as landscaping. Plastic containers
are available in a variety of shapes and sizes, and are reusable after sterilization. Be sure that
containers are suitable for the root growth of the plant in question. Citrus pots are nearly twice as
tall as standard one-gallon containers, and provide adequate depth for some of the native shrubs
and trees that have deeper tap roots. Planting instructions are similar to trays and plant bands,
except that the soil is not as tightly packed into the container.

Potting Mediums for Containerized Plants
For small-scale operations, good potting mixes can be purchased commercially. For larger-scale
greenhouse or nursery operations, bulk potting mediums can be specifically mixed to provide
nutrients and adequate drainage to growing seedlings or transplants. Most plants are extremely
adaptable and can tolerate a wide range of soil conditions. Soil mixes can also be adapted to meet
a plant’s specific nutritional, pH, or water requirements. Most soil mixes are made up of some
combination of sand, vermiculite or perlite, and organic matter. Soil conditioners, such as mill slab
or commercial products, promote soil aeration and water penetration, and also encourage microbial
activity. The soil medium of containers should be a similar kind to the planting site so that roots will
readily grow out into the field soil. For example, if the planting site has heavy clay soil, be sure to
use a heavy soil in containers.
Propagation
Once seeds are collected and cleaned, they are ready to be propagated using 1 of 2 basic systems.
The most common is direct seeding and germination in small containers. The second method
involves dividing or splitting adult plants into numerous transplants, which is generally used for
species with poor seed production and quality, low germination, and long dormancy periods.
Grasses
Seed Propagation
If climate controlled greenhouses are not available, late summer or early spring plantings for
grasses are best because of the harsh summer conditions in South Texas. For general purposes,
grasses are best seeded in flats or trays that
measure 1” x 1” x 3” per cell, and contain about
70–200 cells/tray. If using plant bands, then
a 1” x 3” cell is best. If field planting will be
delayed, then a deeper 1” x 6” heavy grade
paper band is recommended to sustain longer
root growth.
The number of seeds to plant per tray or plant
band cell depends on germination percentages
of the selected species. Because limited
germination data are available for native and
locally collected seed, it may be necessary
to contact your USDA-NRCS Plant Materials
Center or local growers to obtain an estimate
on germination rate. A do-it-yourself approach
can also be taken by planting 1 or 2 flats with
a set number of seeds per cell to estimate
germination percentage. This will avoid wasting
seed in larger plantings. If you cannot obtain
the germination percentage by the above
methods, 10% should be assumed.
Seedling nursery
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HELPFUL TIPS
Because nutritional needs are different for some types of plants, potting mixtures may need to be adapted
to be more acidic or basic. A good potting soil mixture for South Texas natives should contain about 1 part
sandy loam soil, 1 part finely ground Mexican perlite, 1 part composted ground lumber mill “slab” (sold as
“soil conditioner”), and 1 part of thoroughly composted feedlot manure. One ounce of slow release 12-
12-12 fertilizer can also be added before mixing each 4-cubic-foot batch. For acid-loving plants, increase
the proportion of organic matter by 1 part. For plants that require good drainage, such as cacti and succulents,
increase the proportion of sand by 1 part.
About Manure
Many animal waste products can be used for manure, such as from cows, sheep, horses, pigs, chickens,
and bats. However, it is critical that animal waste products be fully composted. The composting process
will kill any weed seeds or harmful pathogens remaining in the manure. Manure that has not fully decomposed,
or “green” manure, can starve young seedlings of important nutrients or accumulate metabolic
products that can “burn” young seedlings. In addition, they can also create odors that may attract scavengers
to dig up the seedlings after transplanting.
Do I add topsoil to my mixture?
There are 2 approaches regarding the use of topsoil in greenhouse potting mixes. The first incorporates
native topsoil because it supplies native microbes, which enhance seedling growth and establishment
and help maintain proper soil pH. In this case, it’s best to use a 1:1 native topsoil to vermiculite soil mix,
along with an encapsulated time-released fertilizer. The use of topsoil may be practical for single-site
restoration projects, but not if seeds are being collected at various locations. The second approach does
not use topsoil in potting mixes, but uses “clean” sterilized soil to minimize the spread of fungus and disease
which might be introduced from outside the greenhouse. Sterilized soil, however, will not contain
the other microbes which serve as deterrents for future pathogen invasion.
Once germination percentages have been established, trays should be planted at 2 live seedlings
per cell; more than 2 seedlings may limit seedling growth. If you have a small seed supply, use
only 1-3 seeds per cell even though not all cells will produce seedlings. It’s much easier to dump
out empty cells or pots than to tediously thin out seedlings that are too dense. It is possible
that more seeds than you expected may germinate; in this case, you may have to spend some
time separating and transplanting seedlings into other trays using tweezers. For some species,
however, thinning may take place naturally.
After seeds are planted, trays should be regularly watered with pure (reverse-osmosis, distilled,
or de-ionized) water or collected rainwater. Avoid using tap water since salt levels in South
Texas can negatively impact seed germination and early growth. The soil should be kept
moist, but not super-saturated in the early stages of growth. After the plants have grown to
3” tall, they can be watered less often, but with a deeper soaking. Plants at this stage will
also be less sensitive to poorer water quality.
After seeding, wait 30–45 days to give the seeds ample time to germinate. Once
seedlings have germinated, allow approximately 2–3 months growing time. If seedlings
begin to demonstrate nutrient deficiency symptoms, a liquid fertilizer such as
Miracle Gro®, seaweed, compost tea, or other compatible brands can
be added with water. Some plants are sensitive to micronutrient levels
such as iron and will require an additional fertilizer application. Plants are
ready for transplanting when they have a well-developed root system that
holds the soil together when the seedlings are pulled from trays.
102
Arizona cottontop (Digitaria californica) seedling

Vegetative Propagation
Plants can be propagated through division
or cuttings from live plants, and this method
is generally used for species with poor
seed production and seed quality. It is
also beneficial when selecting for specific
plant characteristics or when only a few
plants are needed. New plants can be
produced in shorter periods of time, as is
often done in the nursery trade business.
When selecting which plants to propagate,
choose vigorous individuals because new
plants will be identical to the parent plants.
If your goal is to transplant into the wild,
you should propagate from a number of
plants to maintain genetic diversity. If
propagating plants for commercial use or
landscaping, however, genetic replication
is not a problem.
Division is a simple method that can
be used for bunch grasses such a
gulf cordgrass and switchgrass, and
should be done after the onset of active
growth (usually spring), but before seed
production begins. Large clumps can be
extracted from the soil either with a shovel
or with a backhoe. The clumps can then
be divided with a hatchet, quartering
natural cleavage lines into small bunches
that usually consist of 2 to 3 tillers.
Bunches can be planted immediately
using the existing soil that has been
amended with potting or native soil; tops
should be cut back to ¹⁄3 the height of the
plant, or approximately 6”. If bunches
will be planted at a later time, excess soil
should be shaken off and the roots rinsed
in water. Plants should be kept wet to
prevent bunches from drying out until they
are transplanted into the potting medium.
Root material should be trimmed to about
3”, and the top material should be trimmed.
Bunches can then be planted into 1” x 8”
plastic containers that are half filled with
commercial, fine textured soil mix, and
then filled approximately ¾” from the top.
Plant division is a useful technique for propagating native
grasses.
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Cuttings from rhizomatous and stoloniferous plant material such as marshhay cordgrass, seashore
dropseed, and curlymesquite can be similarly propagated. Stolons or rhizomes can be extracted
from the soil and cut into pieces containing at least one node; each node should have an emerging
shoot and root. Pieces are then planted into 1” x 3” or 1” x 6” paper plant bands, pots, or sprigged
directly into the ground. Fertilizer can be used, but be careful to use diluted liquid or slow-release
soil fertilizer to prevent burning new roots. Plants should be ready to transplant in 1-2 months.
Forbs
Seed Propagation
Most forbs are propagated by seed, and are planted and maintained similar to grasses. Because
most forbs have a taproot instead of the fibrous roots of grasses, they should be handled carefully
to avoid bending or causing “J-rooting” of the taproot while transplanting. Smaller forbs can be
germinated in larger trays or flats that measure 1.5” x 2” x 2.5” per cell.
If seeded in large flats, seedlings will be ready to transplant into larger, 6” deep sterilized containers
after the first true leaves appear. Trays should be watered thoroughly to make removal of the
seedlings easier. Seedlings are transplanted by lightly pinching the true leaves between the index
finger and thumb; this will help prevent damage and the introduction of soil borne disease to the
stem. Be careful not to leave the roots exposed to air for any length of time. Certain types of
seedlings produce “rosettes” at the stem base instead of elongated stems. These seedlings should
be transplanted at the same depth as when growing in the trays or flats to avoid covering any basal
growth.
Larger woody forbs and smaller shrubs can also be propagated in trays if cells provide sufficient
room for roots to grow and for the seedling to develop into a healthy transplanting size. If field
transplanting might be delayed, a deeper cell (e.g., 6”) may be best to allow for more root growth.
If cost and space allow, propagating forbs in individual plant bands or containers may be better
than using trays because there is no need to disturb the root system. This is certainly the case for
larger, woody-stemmed forbs. Before transplanting to the field, the plants should have a stable root
system that reaches the bottom of the transplant containers, or spiral around the sides.
Containers with native forbs in greenhouse
104

Vegetative Propagation
Division can be used for forbs such as gayfeather and wild onion, which have tubers and bulbs
that can be separated into several plants. These are propagated the same as with grass divisions.
Generally, divide plants after they have seeded, or early in the spring when leaves first appear.
Shrubs
Seed Propagation
Shrubs require more established root systems before transplanting, so plants should be allowed
to reach a larger size than that required for smaller forbs and grasses. Shrub seeds should be
germinated in 2” wide plant bands between 6”–12” tall, in small 1” x 8” containers, or if space
allows, in citrus pots.
Vegetative Propagation
Woody plant materials or shrubs such as marsh elder, guayacan, and armed saltbush can be
propagated by clipping the 3-6” apical tips from branches with new growth. Containers should be
filled ¾” from the top; a small hole is formed using a pencil or some other small object. The cuttings
are then dipped into a rooting hormone before being inserted into the hole and planted into the 1”
by 8” plastic containers. Gently press the soil around the cutting, covering at least one node.
Cuttings can be covered with airtight domed plastic covers
or with translucent plastic bags, which will increase humidity
and encourage growth. Another technique is to place the
shrub transplants into a container, such as a baby pool,
filled with water about half way up the plant containers.
The holes in the plant containers will allow water to wick
upwards and keep the soil moist. After about 2 months
in the baby pools, transplants can be moved to a shade
house or placed under a tree. They should receive only
top-watering and become acclimated to outside conditions
before being planted in the field. Pricklypear cactus is a
species that can be easily propagated by placing cut pads
in moist soil.
A few shrub species, such as Carolina wolfberry can be
Aquatic plants for transplanting
propagated through division. Procedures are similar to that of bunch grass division.
Grass seedlings in trays
105

Seedling Maintenance
Grasses, forbs, and shrubs
An environment that provides good ventilation and adequate light will minimize disease for growing
seedlings. Maintaining an adequate watering system is also critical. If the surface of the soil in the
container is dry but the remainder is still moist, the seedling does not need to be watered. If left
too long without water, however, the seedling can wilt and die. If possible, avoid leaving water on
the leaves during the night, which can cause disease. Watering during early morning will alleviate
this problem. Make sure irrigation flow is even and gentle to prevent damage and stem breakage
to seedlings.
Many seedlings will respond well to weekly applications of liquid fertilizer such as 20-20-20. Organic
fertilizers such as seaweed and humate-based products also contain important trace elements. Be
careful not to over-fertilize (noted by foliage discoloration or poor condition), and do not fertilize on
hot, sunny days to avoid burning foliage.
Transplanting Native Plants
Grass plants should be trimmed and maintained at about 6” tall until transplanting. Shrub seedlings
should be pruned several times during the growing season while they are still in the nursery, keeping
them about 1’ tall. Prune no more than ¹⁄3 of the plant during each pruning. Pruning encourages
root growth, maintains the proportion of top growth to root mass, and helps discourage herbivory
after the seedlings are transplanted into the field. Before field planting, transplants should have
abundant roots that hold the soil together, but roots should not wind around container. Generally,
the above ground portion should be 1-2 times the root depth. Seedlings should also be moved 2
or 3 times to prevent roots from growing into neighboring containers. Moving containers on the
bench provides an opportunity to identify plants that fail to germinate, and to arrange seedlings so
that larger plants do not shade out smaller ones.
106
Hardening off seedlings before transplanting

If plants have not been field planted by April 1 (in South Texas), greenhouse propagated seedlings
should be acclimated to uncontrolled environmental conditions (or “hardened off”) before they are
transplanted. This can be done using a shade house with a 20% shade cover, and where seedlings
receive only top watering. By the end of the growing season, plants should have been in full sun
for at least a month, and will be ready for transplanting.
Transplanting Plants
Seeds
For small projects, seedlings can be transplanted by
hand using a shovel or trowel. Mechanical transplanters
can be used for larger quantities, which substantially
speed-up the planting
process.
These machines work
best on well-cultivated,
loamy soils. Coarse
textured, sandy soils
may not form an
adequate planting slot,
but with specialized
equipment and moist
soil conditions, these
Mechanical transplanter
slots can be created. Fine textured, heavy clay soils may
also be somewhat challenging to plow, but thorough and
repetitive tilling will provide sufficient depth for planting
transplants.
When seedlings
are ready to be
Transplanting by hand
transplanted,
they are simply pulled out of the cells with the soil intact
and placed in the transplanting device, or planted directly
into the ground. Mechanical
transplanters can also be
adapted to apply water at
each transplant slot before
the plug is transplanted. This
reduces the risk of transplant
shock and facilitates initial
establishment. If using plant
bands, special transplanting
machines are available.
Remember to fully
bury plant bands or else the
paper can wick important
moisture away from the root
system.
Plant band depths; left-wrong, right-correct
107

Once planted, the soil should be firmly packed around the
transplants to prevent moisture loss. If irrigation is available,
seedlings should be watered immediately. In general,
maintenance procedures are the same for transplants as
for a seeded field.
Do not
remove plants from
public sites, unless
authorized!
Shrubs and Trees
Transplanting shrubs and trees can be used to supplement the
diversity of a restoration site or to rescue plants that would otherwise
be destroyed by development or cultivation. When digging wild plants,
make sure the source site has an ample population of the species you are
removing and that only a few plants are taken.
For transplanting shrubs and trees, it is best to target plants when they are less
than 4’ tall. If at all possible, try to obtain plants that are not rooted in rocks; removal may sever
roots or cause too much distress to the plant. To estimate the distance for digging around the
plant, simply measure the crown or spread of the branches; this will be similar to the spread of the
roots underground. While digging, preserve as much soil around the roots as possible. If there
are any broken or extra long roots (excluding the tap root), trim them off with a clean cut. Wrap
the root ball or bare roots with wet burlap as soon as possible, or transfer directly to a large pot
conducive to the size of the root ball. Carry the plant by the root ball (or pot), but not by the stem
or trunk. Transplanting should take place during the early morning before heat induces wilting.
Planting depth should be the same or slightly higher (½” – 1”) than the
depth of the planting container or root ball. In poorly drained clay or
compacted soils, trees and shrubs should be planted 2–4” higher
than the surrounding soil. Also, loosened soil in the bottom of a
hole can settle, causing the plant to be buried too deeply; this
can cause crown rot, to which many of our native trees and
shrubs are susceptible. The ideal width of planting holes is
3x the width of the container or root ball.
Root ball level
or slightly above
existing soil line
Flare
Planting hole Root ball Loosened soil
108
Root ball

Maintenance after Transplanting
Once the plant has been transplanted, trim it back by 1/2 to 2/3 in order to minimize transplant
shock, and keep the soil moist. Transplanting can be relatively costly compared to seeding,
so planned irrigation and weed control are prudent. If water and labor are available,
it is important to irrigate transplants on a regular basis until they are fully established.
Competition for water can limit establishment if other vegetation has not been controlled
in the seedbed before transplanting. Chemicals may be an option for vegetation control
if they can be applied safely over transplants while still effectively controlling the weeds.
Seedlings can be covered or protected by applying herbicide to the targeted weeds with
a hooded sprayer to minimize drift. If chemical control is not an option, mechanically
controlling weed infestations is suggested. In this case, the project design and spacing
of transplants should be considered so that machinery can be operated without lowering
survival and success rate of the seedlings.
Browsing by large and small mammals can be expected, therefore, transplanted
seedlings will need to be protected. Brush species that have physical defenses such
as thorns will have a greater chance of surviving than
seedlings that do not. Protection of seedlings against
browsing may be more expensive and labor intensive,
but will improve seedling establishment. If protection
from browsing is not an option, planting more vulnerable
species in groups surrounded by more browse-resistant
seedlings may help reduce losses.
LITERATURE CITED
Nokes, J. 2001. How to Grow Native Plants of Texas and the Southwest. University of Texas Press, Austin, Texas.
Phillips, H. R. 1985. Growing and Propagating Wild Flowers. The University of North Carolina Press, Chapel Hill and
London.
Arbury, J. B., M. H. Richard, C. Innes, and M. Salmon. 1997. The Complete Book of Plant Propagation. The Taunton
Press, Newtown, Connecticut.
Young, J. A., and C. G. Young. 1986. Collecting, Processing and Germinating Seeds of Wildland Plants. Timber Press,
Inc., Portland, Oregon.
Hopkins, W. G. and M. P. A. Huner, 2002. Introduction to Plant Physiology. Wiley Textbooks, NJ.
http://www.tpwd.state.tx.us/nature/wildscapes/plantsrc.htm
http://www.nativeplantnetwork.org
109

110
Notes

Coastal Prairie, Cameron Co., Texas
MANAGEMENT
PRACTICES AFTER
RESTORATION
111

112

MANAGEMENT PRACTICES
AFTER RESTORATION
The goal of maintaining or improving the quality of a reseeded or restored site should include
management plans for livestock and wildlife, and protection from vehicular traffic, such as ATV’s
or trucks. The integration of follow-up treatments to manage vegetation, reduce erosion, and or
improve wildlife habitat are also important. Treatments might include brush management, prescribed
burning, and the addition of fences and watering facilities to improve grazing distribution. In addition,
proper grazing management of domestic livestock and wildlife will be absolutely necessary to
successfully restore the site.
Important Factors to Consider before Grazing or
Burning a Newly Restored Site
Restoration is a process, and without a planned grazing and prescribed burning program that
considers the fragility of seedling plants, the restoration process can be delayed or negatively
impacted. Many disturbances can negatively impact plant survival and establishment, and may
reduce species diversity. Sometimes, disturbances can produce an environment favorable for
exotic or woody plant invasion. Furthermore, different species have different responses to grazing
or burning. The following are considerations that should be made before you implement any
management strategy:
Be sure you carefully
consider any management
practice that might cause
disturbance on a newly
restored site!
• Develop a plan that considers the ecological site or soil series
description.
• Evaluate the restoration site; determine whether the plants are
well established, are the desired species, and at the appropriate
densities.
• Understand the morphology and reproductive mechanisms of plants on
the restoration site. For example, whether the plants reproduce by seed, or
asexually through tillers or rhizomes, will be important because each has a different
response to grazing and burning.
• Understand the recovery period requirements of species after grazing or burning for
vegetative, root, and reproductive growth.
• Determine the utilization limits for plant species and optimal grazing seasons.
• Determine the types of grazing strategies used by the animals that will use the site, including both
wild and domestic animals, rodents, or insects.
• Estimate the amount of litter or mulch accumulated at the site. This will determine the amount
of organic matter that will contribute to the productivity of the system, and help protect against
erosion.
• Consider the amount of rainfall the site has had in the past 2 years; high rainfall will contribute
to faster recovery because of high soil moisture, but dry soils may delay the process.
• Consider the life cycle (perrenial, annual, biannual) and season of growth of each species;
response to burning or grazing will vary by species.
113

Grazing
The goal of prescribed grazing is to control vegetation harvest through
grazing or browsing animals. The maintenance of plant health and vigor
is critical at this stage of a restoration project. Careful management of
stocking rates and grazing duration are important considerations.
As a rule, recently reseeded or restored rangeland should not be grazed
for a minimum of 2 years. As with all rangeland practices, this period
will vary from region to region based on climatic factors such as annual
rainfall and soil type; in some cases, some newly restored areas may
need to be rested as much as 4 years. Perennial grasses, forbs, and
shrubs need several years to establish root systems and have enough aboveground
size to withstand removal by grazing. Following 2 years of total rest, if
the plant community has become established, the area can be grazed based on a
sound grazing plan. Stocking rates should be based on objectives of the restoration
project. Some examples of obectives could be to reduce litter, control woody vegetation, or
to promote forb growth.
Prescribed Burning
Reasons to burn a restoration site may include improving forage quality, controlling woody plants,
and decreasing litter or increasing forb growth. In areas where increasing shrubs are a concern,
prescribed burns can be used when shrub seedlings are first noted, but only if the area has been
rested sufficiently (> 2 years). If shrubs begin to appear before the area can sustain a prescribed
burn, unwanted plants can be spot-treated with herbicides.
Fire is a natural and integral component of South Texas rangelands. Much like their adaptation
to variable rainfall, many plants, such as bunchgrasses and deeply-rooted forbs, are adapted to
seasonal fires, which historically helped to maintain grasslands. Much like grazing, prescribed
burning on a restored site depends on the plant production and species composition.
Most newly reseeded or restored land should not be burned within the first 2 years after seeding
or planting because of shallow roots and immature plant vigor. Guidelines for prescribed burns on
South Texas rangelands apply to restored sites as well. A successful fire largely depends on the
accumulation of fine fuel to carry the fire.
Seasonal timing for prescribed burning will depend on the management goals for the site, and should
already have been predetermined in the initial plan. Specific timing will be based on environmental
conditions. Fuel loads, air temperature, wind, relative humidity,
and soil moisture are critical factors, and will have to be estimated
before a prescribed burn.
114

Grazing Following Controlled Burns
Factors affecting Length of Post-fire rest before grazing:
• Ecosystem type and ecological condition before burning.
• Vigor of vegetation before the fire.
• Season of the fire.
• Classic factors affecting growing season conditions, including
temperature and precipitation, before and following burning.
Grazing duration and stocking
rates may need to be adjusted
throughout the year!
• The management objectives for the area; for example, the length of postfire rest
may vary significantly for the same area depending upon whether the area is
being managed primarily for livestock, wildlife, recreation, or a combination of
these activities.
• When determining stocking rates for burned sites, it is important to consider
that livestock and wildlife will concentrate on burned areas because of new and
palatable plant growth.
Wildlflowers and mixed shrub-grassland, Kleberg Co., TX
LITERATURE CITED
Rangeland Health: New Methods to Classify, Inventory, and Monitor Rangelands. By the Committee on Rangeland
Classification, Board on Agriculture, National Research Council, National Academy Press, Washington, DC 1994
Standards and Guidelines for Healthy Rangelands, Bureau of Land Management: BLM-UT-GI-97-001-4000; U.S. Department
of Interior Bureau of Land Management, Utah State Office 1997
115

116
Notes

Sunrise, Webb Co., TX
APPENDICES
117

APPENDIX A
Glossary of Plant Basics
Much of the information published in this appendix was adapted from the Native Plant Revegetation
Guide for Colorado - Caring for the Land Series Volume III, Colorado Natural Areas Program,
Colorado State Parks, and Colorado Department of Natural Resources, 1998.
The Plant Kingdom is extremely diverse, and contains over 200,000 species. Taxonomists combine
these species into a variety of groups and sub-groups on the basis of shared characteristics. The
majority of plants discussed in this guide belong to the large category of plants that reproduce by
means of seeds (i.e., spermatophytes).
Two broadly defined groups of seed plants are gymnosperms and angiosperms.
Gymnosperms
• Woody plants with needle-like or scale-like (imbricate, overlapping) leaves.
• Do not produce flowers.
• Fruits are cones or berry-like cones (the blue “berries” on junipers are modified cones).
• Includes pines, firs, Douglas-fir, spruces, junipers, and mormon tea from the primitive family of
Ephedraceae.
Angiosperms
• Plants that produce flowers.
• Include species with obvious flowers such as ceniza and sunflowers (Helianthus spp.), as well
as those that are not so obvious, like grasses, bulrushes, and sedges.
Deciduous
• Produce leaves through the growing season, and may drop leaves in the winter. Some plants,
like whitebrush, may drop their leaves during drought.
• Pecan, hackberry, and honey-mesquite are examples of deciduous species.
• Deciduous plants in South Texas may retain their leaves during the winter due to our warm
climate; deciduous plants in most areas of the country are bare during the winter.
Evergreen
• Produce green leaves throughout the winter.
• Guayacan, live oak, anacua, and evergreen sumac are examples of evergreen species.
Both gymnosperms and angiosperms may be either deciduous or evergreen.
Gymnosperms and angiosperms can be further categorized by life cycle, growth form,
reproductive strategy, maturity rate, and other descriptive terms.
Life Cycles
This manual uses only the general life cycle terms annual, biennial, and perennial.
118

Annual plants live for only one growing season. For annual species, the emphasis of growth
is placed on production of seeds. Above ground growth is rapid because the plant must flower
and produce seed before it dies at the end of the season. As a consequence, annual species can
be recognized by their relatively small root systems. Often annual plants are seeded at locations
that need to be vegetated quickly, such as highly erosive slopes. While annuals grow quickly
and provide cover, they do not provide a long-term solution to the challenge of revegetation. The
seed that they produce, however, may be stored in the soil. These plants will continue to appear,
competing for nutrients, water and sunlight, long after the area has been planted with species that
are more permanent. Many weeds and crop plants are annuals.
Biennial plants live for 2 years. During the first year they typically produce rosettes of basal
leaves and store energy. They send up flowering stalks the second year, produce seeds, and die.
Native biennials are relatively uncommon, however, many invasive weeds are biennials.
Perennial plants live for many years. Normally they do not flower or reproduce until they are
several years old. Perennials can afford to direct more energy to long-term establishment and
typically have more extensive root systems. These species are usually the most desirable for
native plantings.
Growth Forms
♦ Woody plants form stems composed of lignin that mature to wood; these include
trees, shrubs, and woody vines. They do not die back to ground level each winter as
do forbs, grasses, and grass-like plants. On woody plants, buds are produced above
ground level, giving these plants a “head start” to larger stature each spring.
▪
▪
▪
Trees are woody perennial plants that typically have a single main stem
or trunk and radiating branches on the upper portion of the plant. They
usually stand over 13 feet tall at maturity.
Shrubs are woody perennials lacking a main stem, instead having several
to many branches arising at ground level. They are usually less than 10 to
13 feet tall.
Vines are climbing or trailing plants with long, slender stems that trail on
the ground, or climb around a fixed object.
♦
Herbaceous plants have little or no woody tissue
▪ Graminoids include grasses and grass-like plants with parallel venation
such as sedges and rushes.
♦ Grasses have jointed, hollow stems and clusters of small
membranous flowers arranged in spikelets.
♦ Sedges resemble grasses but have solid stems which are often
triangular.
♦ Rushes resemble grasses but have round, pliant, hollow, or pithy
stems lacking joints.
▪ Forbs are broad-leaved herbaceous plants that die back to the ground
each year and are collectively known as wildflowers.
119

Reproductive Strategies
Seed reproduction is found in all groups of gymnosperms and angiosperms. Seeds may be spread
by wind, water, animals, or humans. Some seeds may require special treatment or conditions
such as fire, freezing temperatures, or passage through the digestive tract of animals in order to
germinate.
Strategies
Applications
Rhizomatous forbs and
graminoids have underground
stems and branches that
spread in linear patterns.
New plants grow from these
rhizomes.
Rhizomatous forbs and
graminoids are ideal for erosion
control because their extensive
networks of underground
stems retain soil. E.g., prairie
acacia and switchgrass.
Bunch-forming forbs and
graminoids grow in clusters
and appear as round tufts on
the landscape. New sprouts
(tillers) grow only from the
base of the plant, increasing
the width of the bunch over
time.
Stoloniferous forbs and
graminoids have above ground
stems that spread, root, and
sprout new individuals.
Vegetative reproduction
Many shrubs, grasses, and a few trees spread by root
shoots or points on the roots from which new stems
or trunks can emerge. Most forbs and graminoids
reproduce by seed; however, many perennials have
developed additional strategies for propagation.
Bunch-forming plants
are valuable for ground
nesting bird habitat (e.g.,
little bluestem, tanglehead,
paspalums, multiflowered false
rhodesgrass).
Stoloniferous graminoids are
useful for heavy-use areas
and erosion control because
they form durable sods (e.g.,
curly mesquite, buffalograss,
and shortspike windmillgrass).
HELPFUL TIPS
Planting a mixture of both warm and cool
season plants provides robust year-long
cover and visual interest from spring to fall.
Seasonal Growth
Seasonal growth patterns are important considerations, especially for grass plantings. Two general
patterns are recognized – cool season and warm season.
Cool season graminoids begin their growth in late winter and early spring and bloom in spring
or early summer. They may enter dormancy during summer heat and resume growth or even bloom
again in the fall if adequate moisture is available (e.g., Canada wildrye and Texas wintergrass).
Warm season graminoids begin their growth in early spring or summer and usually bloom in
late summer or early fall, and may bloom several times during one growing season. These plants
then usually enter dormancy with the onset of winter (e.g., pink pappusgrass, Plains bristlegrass).
120

Seed Coat
A seed coat is the protective outer covering of the seed. It protects the embryo from injury and from
drying out. The thickness of seed coats is important to consider for propagation efforts.
Hard seeds have thick seed coats, and may require some scarification for germination to occur.
Many shrubs have hard seeds, such as those found in the the Fabaceae family (legumes) (e.g.,
mesquite, blackbrush, and mountain laurel).
Soft seeds have thin seed coats, and will likely germinate easily on their own. These include the
majority of the graminoids and many forbs.
121

APPENDIX B
Suggested Plant Species for
Use in Restoration, Listed by Communities
Vegetation associations are those defined by McLendon (1991). Species listed are those of
importance for wildlife and livestock, and have shown potential for successful use in revegetation/
restoration projects. Plants with an * are those that have been released for commercial production
by South Texas Natives or the USDA-NRCS E. Kika de la Garza Plant Materials Center as of
January 1, 2008.
Community
Grasslands
Little Bluestem-Trichloris Association
Little bluestem (Schizachyrium scoparium)
Multiflowered false rhodesgrass (Trichloris pluriflora)
Silver bluestem (Bothriochloa laguroides)
Big bluestem (Andropogon gerardii)
Plains bristlegrass (Setaria leucopila)*
Whiplash pappusgrass (Pappophorum vaginatum)
Sideoats grama (Boutleoua curtipendula)
Slender grama (Bouteloua repens)*
Hairy grama (Bouteloua hirsuta)*
Texas grama (Boutleoua ridigiseta)*
Buffalograss (Buchloe dactyloides)
Curly mesquite (Hilaria belangeri)
Hooded windmillgrass (Chloris cucullata)*
Shortspike windmillgrass (Chloris x subdolistachya)*
Plains lovegrass (Eragrostis intermedia)
Texas wintergrass (Stipa leucotricha)
Orange zexmenia (Wedelia hispida)*
Awnless bush sunflower (Simsia calva)
Engelmann daisy (Engelmannia pinnatifida)
Dayflower (Commelina erecta)
Western ragweed (Ambrosia psilostachya)
Croton (Croton spp.)
Gayfeather (Liatris spp.)
Sunflower (Helianthus spp.)
Plantain (Plantago spp.)*
Bundleflower (Desmanthus virgatus)
Deer pea vetch (Vicia ludoviciana)
Prickly pear (Opuntia spp.)
Prairie acacia (Acacia angustissima)
Clammyweed (Polanisia erosa)*
122

Community
Seacoast bluestem-balsamscale Association
Seacoast bluestem (Schizachyrium littoralis)
Panamerican balsamscale (Elyonurus tripsacoides)
Tanglehead (Heteropogon contortus)
Brownseed paspalum (Paspalum plicatulum)
Gulfdune paspalum (Paspalum monostachyum)
Marshay cordgrass (Spartina patens)
Thin paspalum (Paspalum setaceum)
Switchgrass (Panicum virgatum)
Plains bristlegrass (Setaria leucopila)*
Slender grama (Bouteloua repens)*
Hairy grama (Bouteloua hirsuta)*
Threeawn (Aristida spp.)
Sand dropseed (Sporobolus cryptandrus)
Texasgrass (Vaseyochloa multinervosa)
Crinkleawn (Trachypogon secundus)
Big bluestem (Andropogon gerardii)
Yellow Indiangrass (Sorghastrum nutans)
Partridge pea (Chamaecrista fasciculata)
Lazy daisy (Aphanostepus spp.)
Dayflower (Commelina erecta)
Sunflower (Helianthus spp.)
Snoutbean (Rhynchosia spp.)
Orange zexmenia (Wedelia hispida)*
Awnless bushsunflower (Simsia calva)
Croton (Croton spp.)
Indian blanket (Gallardia spp.)
Dalea (Dalea spp.)
Plantain (Plantago spp.)*
Prickly pear (Opuntia spp.)
Woodlands
Hackberry-huisache Association
Sugar hackberry (Celtis laevigata)
Huisache (Acacia smallii)
Canada wildrye (Elymus canadensis)*
Virgina wildrye (Elymus virginianus)
Bristlegrass (Setaria spp.)*
Multiflowered false rhodesgrass (Trichloris pluriflora)
Brownseed paspalum (Paspalum plicatulum)
Eastern gamagrass (Tripsacum dactyloides)
Texas wintergrass (Stipa leucotricha)
Mistflower (Eupatorium odoratum)
123

Community
Hackberry-huisache Association (cont’d)
Turks cap (Malvaviscus arboreus)
Pigeonberry (Rivina humilus)
Texas sage (Salivia coccinea)
Anaqua (Ehretia anacua)
Ebony (Pithecellobium ebano)
Live Oak-Post Oak Association
Live oak (Quercus virginanus)
Post oak (Quercus stellata)
Seacoast bluestem (Schizachryium littoralis)
Little bluestem (Schizachyrium scoparium)
Texasgrass (Vaseyochloa multinervosa)
Slender grama (Bouteloua repens)*
Mistflower (Eupatorium odoratum)
Turks cap (Malvaviscus arboreus)
Texas sage (Salvia coccinea)
Hooded windmillgrass (Chloris cucullata)*
Hairy grama (Boutleoua hirsuta)*
Sand dropseed (Sporobolus cryptandrus)
Multiflowered false rhodesgrass (Trichloris pluriflora)
Awnless bushsunflower (Simsia calva)
Shrublands
Mesquite-Granjeño Association
Little bluestem (Schizachyrium scoparium)
Silver bluestem (Bothriochloa laguroides)
Cane bluestem (Bothriochloa barbinodis)
Multiflowered false rhodesgrass (Trichloris pluriflora)
False rhodesgrass (Trichloris crinita)*
Plains bristlegrass (Setaria leucopila)*
Arizona cottontop (Digitaria californica)*
Pink pappusgrass (Pappophorum bicolor)
Whiplash pappusgrass (Pappophorum vaginatum)
Vine-mesquite (Panicum obtusum)
Hooded windmillgrass (Chloris cucullata)*
Shortspike windmillgrass (Chloris x subdolistachya)*
Hairy grama (Boutleoua hirsuta)*
Orange zexmenia (Wedelia hispida)*
Awnless bush sunflower (Simsia calva)
Bundleflower (Desmanthus virgatus)
Prickly pear (Opuntia spp.)
124

Community
Mesquite-Granjeño Association (cont’d)
Spiny hackberry (Celtis pallida)
Brasil (Condalia hookerani)
Hogplum (Columbrina texana)
Coma (Bumelia spp.)
Prairie acacia (Acacia angustissima)
Clammyweed (Polanisia erosa)*
Plantain spp. (Plantago spp.)*
Huisache-Prickly Pear Association
Silver bluestem (Bothriochloa laguroides)
Alkali sacaton (Sporobolus airoides)
Big sacaton (Sporobolus wrightii)*
Gulf cordgrass (Spartina spartinae)
Knotroot bristlegrass (Setaria geniculata)
Vine mesquite (Panicum obtusum)
Multiflowered false rhodesgrass (Trichloris pluriflora)
Mesquite-Prickly Pear Association
Curly mesquite (Hilaria belangeri)
Alkali sacaton (Sporobolus airoides)
Red grama (Bouteloua trifida)
Texas grama (Bouteloua rigidiseta)*
Threeawn (Aristida spp.)
Whorled dropseed (Sporobolus phramidatus)
Tobosa (Hiliaria mutica)
Plains bristlegrass (Setaria leucopila)*
Pink pappusgrass (Pappophorum bicolor)
Whiplash pappusgrass (Pappophorum vaginatum)
Multiflowered false rhodesgrass (Trichloris pluriflora)
False rhodesgrass (Trichloris crinita)*
Arizona cottontop (Digitaria californica)*
Gulf cordgrass (Spartina spartinae)
Deer pea vetch (Vicia ludoviciana)*
Bundleflower (Desmanthus virgatus)
Prickly pear (Opuntia spp.)
Guajillo-Cenizo Association
Guajillo (Acacia berlandieri)
Cenizo (Leucophyllum frutescens)
Blackbrush acacia (Acacia rigidula)
Texas kidneywood (Eysenhardtia texana)
125

Community
Guajillo-Cenizo Association (cont’d)
Spanish dagger (Yucca treculeana)
Prickly pear (Opuntia spp.)
Mountain laurel (Sophora secundiflora)
Guayacan (Porlieria angustifolium)
Shrubby blue sage (Salvia balletiflora)
Leatherstem (Jatropha diocia)
Curly mesquite (Hilaria belangeria)
Texas grama (Bouteloua rigidiseta)*
Slender grama (Bouteloua repens)*
Sideoats grama (Bouteloua curtipendula)
Slim tridens (Tridens muticus)
Threeawn (Aristida spp.)
Plains bristlegrass (Setaria leucopila)*
Hooded windmillgrass (Chloris cucullata)*
Silver bluestem (Bothriochloa laguroides)
Cane bluestem (Bothriochloa barbinodis)
Pink pappusgrass (Pappophorum bicolor)
Arizona cottontop (Digitaria californica)*
Orange zexmenia (Wedelia hispida)*
Awnless bushsunflower (Simsia calva)
Blackbrush-Twisted Acacia Association
Blackbrush acacia (Acacia rigidula)
Twisted acacia (Acacia tortuosa)
Texas kidneywood (Eysenhardtia texana)
Spanish dagger (Yucca treculeana)
Prickly pear (Opuntia spp.)
Guayacan (Porlieria angustifolium)
Curly mesquite (Hilaria belangeria)
Texas grama (Bouteloua rigidiseta)*
Slender grama (Bouteloua repens)*
Sideoats grama (Bouteloua curtipendula)
Slim tridens (Tridens muticus)
Threeawn (Aristida spp.)
Plains bristlegrass (Setaria leucopila)*
Hooded windmillgrass (Chloris cucullata)*
Silver bluestem (Bothriochloa laguroides)
Cane bluestem (Bothriochloa barbinodis)
Pink pappusgrass (Pappophorum bicolor)
Arizona cottontop (Digitaria californica)*
Orange zexmenia (Wedelia hispida)*
Awnless bushsunflower (Simsia calva)
126

Community
Creosotebush-Prickly Pear Association
Creosotebush (Larrea divaricata)
Prickly pear (Opuntia spp.)
Saltbush (Atriplex spp.)
Whiplash pappusgrass (Pappophorum vaginatum)
Arizona cottontop (Digitaria californica)*
Curly mesquite (Hilaria belangeria)
Slim tridens (Tridens muticus)
Plains bristlegrass (Setaria leucopila)*
LITERATURE CITED
McClendon, T. 1991. Preliminary description of the vegetation of South Texas exclusive of coastal saline
zones. Texas Journal of Science 43:13-32.
127

APPENDIX C
Selected Plant Profiles
Seeding rates are for planting pure stands. Seeding rates should be adjusted accordingly for
planting in seed mixes. Germination rate is the average % of that observed within the species.
Agarito (Mahonia trifoliata)
Agarito is an evergreen shrub found throughout the Rio Grande Plains. It grows on a variety of
sites but most frequently on gravelly hillsides as part of a mixed brush community. It is an excellent
wildlife plant, as many species of wildlife readily consume the red fruit. Agarito seed should be
cleaned upon collection. Each berry contains several seed. Seeds should be cold scarified to
break embryo dormancy. Natural germination of agarito is low. Agarito is difficult to grow from
cuttings.
Germination Seeding rate Seeds/pound Planting depth Seed maturity
May-June
Agarito
Arizona cottontop (Digitaria californica)
Arizona cottontop is a warm season perennial bunchgrass found in many portions of South Texas.
Plants have white cotton-like seedheads 2-5 inches long. It is considered good forage for cattle.
Arizona cottontop has limited value to wildlife as cover. It is found in a variety of soil types and
commonly grows in areas protected from grazing by dense brush canopies or those areas excluded
from grazing. “LaSalle” is a South Texas release of this species that is now available for planting.
Germination
Seeding rate
Seeds/pound
Planting depth
Seed maturity
47-68%
1-2 lbs PLS/ac.
693,611
≤ 0.25”
April-Nov.
Arizona cottontop
Awnless bush sunflower (Simsia calva)
Awnless bush sunflower is a perennial forb found in South Texas. It is commonly found on sandy
loam sites or on gravelly soils on caliche ridges. Awnless bush sunflower blooms and produces
seed throughout the year following rainfall. Many insects utilize this plant’s attractive flowers.
White-tailed deer are known to consume the leaves. It should be planted by broadcasting. Solid,
cultivated blocks will produce 119 pounds per acre.
Germination
Seeding rate
Seeds/pound
Planting depth
Seed maturity
7-36%
0.15 lbs PLS/ac.
237,593
≤ 0.5”
May-Nov.
Awnless bush sunflower
Big bluestem (Andropogon gerardii var. gerardii)
Big bluestem is a warm season, perennial bunchgrass found on the coastal prairies and in the South
Texas Sand Sheet. It is considered a climax co-dominant, but has been extirpated by grazing from
much of its former native range. Big bluestem also grows throughout the tallgrass prairie region
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of the Great Plains where it is also considered a climax co-dominant. It is usually found in deep
sands of the South Texas Sand Sheet, but occasionally occurs in clay soils in eastern South Texas.
Big bluestem is favored forage of livestock and occurs only in areas that have been deferred or
protected from improper grazing. Wildlife value for big bluestem is fair, it provides good nesting
cover as well as perching and singing sites for grassland birds. Big bluestem responds well to
prescribed fire. It spreads by seed and by underground rhizomes. In South Texas it initiates growth
in early spring and seed heads begin to emerge in mid summer, with seed maturity occurring in
late summer or early fall. Seed heads commonly reach a height 8-10’ tall. Relict stands of big
bluestem should be conserved and protected. Highway right-of-ways are often a good source for
seed collection.
Germination
Seeding rate
Seeds/pound
Planting depth
Seed maturity
2-37%
7-10 lbs PLS/ac.
130,000
0.25-.50”
Sept.-Oct.
Big bluestem
Big sacaton (Sporobolus wrightii)
Big sacaton is a warm season grass that forms dense clumps. It is a coarse, upright bunch grass
that can grow from 3 to 8 ft. tall. Big sacaton is primarily adapted to heavier textured soils in areas
west of the Piney woods. Big sacaton is tolerant of highly alkaline and saline soil. It can tolerate
poorly drained soils and seasonally flooded areas. It is also adapted to dry, rocky draws of West
Texas. Big sacaton may be used in pure stands or as part of a rangeland seeding mix for the highly
alkaline soils of western Texas. It is useful for revegetating saline soils throughout south and west
Texas. It performs well as a grass hedge terrace or windstrip for erosion control. It helps stabilize
watershed structures, stream banks and flood plain areas. It is also useful for wildlife cover. Seedbed
preparation should begin well in advance of planting. Planting can be scheduled for early spring or
where there are minimal cool season weeds, it can also be planted in the fall. Establish a clean,
weed-free seedbed by either tillage or herbicides. The seed can be drilled or broadcast. Plants can
also be grown in small paper containers and then transplanted for establishment of grass hedges
and wind barriers. “Falfurrias” is a South Texas release that is now available for planting.
Germination
Seeding rate
Seeds/pound
Planting depth
Seed maturity
58%
0.5-1 lb PLS/ac.
1,400,000
≤ 0.25”
May-Oct.
Big sacaton
Brasil (Condalia hookeri)
Brasil is an evergreen shrub or small tree commonly found in South Texas. It produces a small
black fruit that is consumed by many mammals and a variety of birds. Deer also browse the leaves.
The seeds should be cleaned from the fruit following collection, if the seed is to be stored. Freshly
collected seeds planted immediately after collection will readily germinate. Seeds stored after
collection should be cold scarified to break dormancy. It can also have favorable germination when
freshly collected without any cleaning. Brasil can also be propagated by semi-hardwood cuttings.
Germination
6-40%
Seeding rate Seeds/pound Planting depth Seed maturity
June-Aug.
Brasil
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Brownseed paspalum (Paspalum plicatulum)
Brownseed paspalum is a warm season bunchgrass that grows on sand and sandy loam soils in
South Texas. Plants from moist, sandy soils usually have very pronounced rhizomes. This grass is
considered a climax dominant on sandy upland sites. It provides good to excellent nesting cover
for bobwhite quail, as well as a hard seed that is consumed by quail and turkey. In addition, it
provides fair forage for livestock. Seeds are easily collected because of the height of the seed head
above the foliage. Seeds mature throughout the year depending on rainfall. However, it is most
common for seeds to reach maturity in the late spring and early to late fall.
Germination
Seeding rate
Seeds/pound
Planting depth
Seed maturity
20-40%
543,936
1/2”
April-Nov.
Brownseed paspalum
Buffalograss (Buchloe dactyloides)
Buffalograss is a warm season, perennial grass common on heavy soils throughout South Texas.
Buffalograss is considered good forage for livestock. It grows as a thick turf, covering the ground
by means of many stolons. It also produces seed, but germination is usually very low. It can be
propagated by collection of stolons, and planting individual nodes on a potting or plug medium. This
species is frequently used as a lawn or turf grass in landscaping and golf courses. Buffalograss is
difficult to grow from seed.
Germination
Seeding rate
Seeds/pound
Planting depth
Seed maturity
0-1%
0.5-6 lbs PLS/ac
40,000 (burs)
> 1/4”
May-Nov.
Buffalograss
Coma (Bumelia celastrina)
Coma is a warm season, shrub or small tree common to South Texas. Coma grows in a variety
of soil types throughout the region, usually in small mottes with numerous trees of varying sizes.
White-tailed deer frequently browse the leaves and a variety of birds and other mammals readily
consume the fruit. Fruits are produced from early to mid summer. The seeds should be cleaned
from the fruit upon collection. Germination can be increased by acid or cold scarification to break
the hard seed coat and dormancy. Seeds planted soon after collection may easily germinate
without treatment.
Germination Seeding rate Seeds/pound Planting depth Seed maturity
May-June
Coma
Common curlymesquite (Hilaria belangeri)
Common curlymesquite is a stoloniferous, warm season, perennial grass that grows throughout
South Texas. It is common on calcareous and alkaline soils in the Rio Grande Plain. Where
present, curly mesquite can be the dominant grass, forming large bunches on the landscape. Curly
mesquite provides good forage for livestock. This grass can be propagated by division of stolons.
Curly mesquite has potential uses as a turf grass, and for use in controlling erosion. Propagation
from seed is extremely difficult.
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Germination
≤ 3%
Seeding rate Seeds/pound Planting depth Seed maturity
May-Nov.
Common curlymesquite
False rhodesgrass (Trichloris crinita)
False rhodesgrass is a warm season, perrenial bunchgrass found on clay, clay loam, and tight
sandy loam range sites in eastern south Texas. This grass has excellent forage value to livestock
and wildlife and provides good nesting cover to ground nesting birds. “Kinney” false rhodesgrass
is a select release of this species that is commercially available for planting in South Texas.
Germination
Seeding rate
Seeds/pound
Planting depth
Seed maturity
20-60%
1 lb PLS/ac
2,451,892
1/4-1/8”
June-Nov.
False rhodesgrass
Golden dalea (Dalea aurea)
Golden dalea is a warm season, perennial legume found in the sandy prairies of South Texas. It is
rated as a fair to good forage for wildlife (white-tailed deer) and fair forage for cattle. Golden dalea
is usually found as single plants scattered within a landscape. It produces abundant seed in mid
summer. Germination can be improved by scarification after removal from the pod. Some daleas
can be grown from cuttings. Cuttings should be made from old-growth near the base of the plant.
Germination
Seeding rate
Seeds/pound
Planting depth
Seed maturity
2-70%
128,732
May-Nov.
Golden dalea
Hairy grama (Bouteloua hirsuta)
Hairy grama is a common perrenial native grass found in sand, sandy loam and gravelly soils in
South Texas. It is a component of many range sites and habitat types. Hairy grama has fair forage
value. Hairy grama is an excellent choice for highway right-of-way plantings on sandy or gravelly
soils in South Texas. “Chaparral” is a South Texas release of this species available for planting.
Germination
Seeding rate
Seeds/pound
Planting depth
Seed maturity
33%
8 lbs PLS/ac
116,300
0.25”
March-Oct.
Hairy grama
Halls panicum (Panicum hallii var. hallii & fillipes)
Halls panicum is a warm season grass common to South Texas. It grows on a variety of soil
types. Determination of variety hallii and fillipes is difficult and some integration between the two
is present in South Texas. Halls panicum is fair livestock forage and has limited value to wildlife.
Birds occasionally eat the seed. Collection can be done by hand or with seed strippers, but care
should be taken to harvest only mature seed, as maturity varies within plants and seed heads. It
has value to restoration efforts because it is a common component of many different successional
stages, and range sites in South Texas. Hall’s panicum has indeterminant seed maturity, making
harvest and seed production difficult.
131

Germination
Seeding rate
Seeds/pound
Planting depth
Seed maturity
10-50%
1 lb PLS/ac.
855,566
≤ 0.25”
May-Nov.
Halls panicum
Hooded windmillgrass (Chloris cucullata)
Hooded windmillgrass is a warm season, tufted perennial grass found throughout South Texas on
a variety of soil types and range sites. Hooded windmillgrass is fair forage for livestock. A high
degree of hybridization with other windmillgrasses causes variation in plant characteristics. Hooded
windmillgrass has potential in habitat restoration. It is an abundant seed producer; seed heads
grow and produce seed rapidly after rainfall events, thus producing seed several times per year
in wet years. Germination is high for hooded windmillgrass. This species can easily be harvested
using conventional harvesting equipment. Hooded windmillgrass’ low successional stage in most
vegetation communities makes it an excellent choice for early establishment in restoration efforts.
“Mariah” is a South Texas release of this species that is now available for planting.
Germination
Seeding rate
Seeds/pound
Planting depth
Seed maturity
≤ 97%
1/4 - 1/2 lbs
PLS/ac.
2,370,654
0.10-0.25”
Feb.-Nov.
Hooded windmillgrass
Lotebush (Ziziphus obtusifolia var. obtusifolia)
Lotebush is a native deciduous shrub common to South Texas. It is occasionally browsed by
white-tailed deer and the blue berries are eaten by many species of wildlife. Fruits have 2 seeds
per stone, and germination can be improved by removal of the seeds from the fruit, and cold
scarification. Seeds are produced in early summer. Lotebush seeds readily germinate immediately
after cleaning and collection.
Germination
95%
Seeding rate Seeds/pound Planting depth Seed maturity
May-June
Lotebush
Mistflower (Conoclinium spp.)
A variety of mistflowers grow in South Texas. Mistflowers are perennial, warm season forbs that
usually grow underneath the canopies of trees and shrubs, although some also grow in open areas
in full sun. They are browsed by white-tailed deer and livestock, and are important butterfly nectar
plants. Mistflowers produce seed throughout the summer and fall. They make nice landscape
plants and attract butterflies when in bloom. Mistflower seeds easily germinate if temperatures are
between 68-86º F. They can also be propagated vegetatively from semi-hardwood and softwood
cuttings. Cuttings should be treated with rooting hormone.
Germination
Seeding rate
Seeds/pound
Planting depth
Seed maturity
50-70%
1,450,496
July-Nov.
Mistflower
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Multiflowered false rhodesgrass (Chloris pluriflora)
Commonly called 4-flower trichloris, this grass occurs throughout South Texas. It is a warm season,
perennial bunchgrass and provides excellent forage for livestock and good cover for wildlife. It is
commonly found on silty, clayey, and sandy loam soils. Multiflowered false rhodesgrass is an
excellent seed producer. As much as 53.5 pounds of seed can be produced per acre in nonirrigated
plots. Seed is produced many times per year and plants can have as many as 47 seed
heads per plant. This grass has high restoration potential because of its abundant seed production
and good germination. A hindrance to restoration efforts is the small, almost weightless seeds,
which are difficult to plant. Seeds should be coated or drilled/broadcasted with a carrier substance
in order to facilitate uniform planting. Planting should take place in late winter or early spring on a
clean, weed free seedbed.
Germination
Seeding rate
Seeds/pound
Planting depth
Seed maturity
20-90%
1 lb PLS/acre
2,145,950
≤ 0.25”
May-Nov.
Multiflowered false rhodesgrass
Orange zexmenia (Wedelia hispida)
Orange zexmenia is a warm season perennial forb common across South Texas. The leaves are
eaten by white-tailed deer and occasionally by cattle, and birds commonly eat the seeds. Orange
zexmenia grows in small clumps and is often found in the protection of small shrubs. Seed can be
produced in dryland settings at a rate of 60 pounds per acre. Seed should be broadcasted in late
winter or early fall on a clean, weed free seedbed. It can also be propagated by semi-hardwood
and softwood cuttings. Transplants or cuttings should be transplanted to the field in early to mid
spring.
Germination
Seeding rate
Seeds/pound
Planting depth
Seed maturity
8-62%
140,520
0.25-0.50”
May-Nov.
Orange zexmenia
Partridge Pea (Chamaecrista fasciculata)
Partridge pea is a long lived, annual legume common to the sandy soils of South Texas. It can be
very common on overgrazed or highly disturbed range sites, but also grows in high successional
areas as well. This species responds well to disturbance, high densities of the plant are usually
present following fire, heavy grazing or mechanical treatment. Partridge pea is considered an
excellent food plant for bobwhite quail and other granivorous birds. It blooms throughout the
summer, but can be most apparent in late summer/early fall after tropical storms. The seeds are
produced in legumes which split upon maturity. Solid plantings can produce an average of 550
pounds of seed per acre. Seeds can be drilled or broadcasted.
Germination
Seeding rate
Seeds/pound
Planting depth
Seed maturity
2-15%
2-10 lbs/acre*
65,000
≤ 0.25”
May-Nov.
Partidge pea
* 2 lbs pls/acre for rangeland seedings, 3 lbs pls/acre for rows, and 10 lbs pls/acre for food plots
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Pink pappusgrass (Pappophorum bicolor)
Pink pappusgrass is a warm season, perennial bunchgrass found in a variety of South Texas
habitats. It is fair forage for livestock. Pink pappusgrass produces a large seedhead that is pinkishpurplish
when immature and turns straw colored when mature. Mature seed heads remain upright,
hold seed well, and are not restricted by foliage. It can be harvested mechanically or by hand.
Germination
Seeding rate
Seeds/pound
Planting depth
Seed maturity
8-30%
3 lbs PLS/ac
398,886
1/4”
May-Nov.
Pink pappusgrass
Plains bristlegrass (Setaria leucopila & vulpiseta)
Plains bristlegrass is a warm season perennial grass found throughout South Texas. It is found in
a variety of soil types and successional stages, and can be especially prevalent on disturbed soils
in the Rio Grande Plain. Identification of the specific species of bristlegrass is difficult to determine
since hybridization likely occurs. Plains bristlegrass is considered good forage for cattle, and
produces seed that is eaten by bobwhite quail and other birds. It produces seed several times per
year. Germination varies because of poor seed fill. Plains bristlegrass should be planted in fall, late
winter, or early spring. Production plots produce 214-369 pounds per acre dependant on seed size.
“Catarina” is a South Texas release of this species that is now available for planting.
Germination
Seeding rate
Seeds/pound
Planting depth
Seed maturity
10-40%
2 lbs PLS/ac.
1,300,000
0.50”
April-Nov.
Plains bristlegrass
Plantains (Plantago spp.)
Plantains (also known as tallow weed) are found across South Texas in a variety of soil types.
Plantains are cool season annual forbs. They are considered good forage for livestock and wildlife.
The seeds are eaten by a variety of game and non-game birds. Seed matures following rainfall and
can be hand harvested by stripping the spiklets of seed using the thumb and forefinger.
Germination
33%
Seeding rate Seeds/pound Planting depth
< 1/4”
Seed maturity
April-July
Plantains
Prairie acacia (Acacia angustissima var. texensis)
Prairie acacia is a warm season perennial legume found in South Texas. It occurs on a variety of
soil types from heavy clays of Eastern South Texas to red sandy loams of the Central Rio Grande
Plain. Remnant stands still remain common on the latter. Large stands of prairie acacia are
present only in areas that are protected from livestock grazing. It grows in colonies, wherein the
majority of reproduction is facilitated by rhizomes. Prairie acacia is considered excellent forage for
livestock and wildlife, and the seeds are known to be eaten by granivorous birds. Generally flowers
and seed are produced following rainfall in the spring through late fall in South Texas. The seed
matures approximately 60 days after flowering. Prairie acacia can also be easily grown from stem
and root cuttings. Stem cuttings should be collected from vigorous plants and placed immediately
on ice or in water (wilting will occur rapidly). Cuttings should be coated in rooting hormone and
134

placed in a firm, well watered potting mixture or soil. Root cuttings can be taken by pulling or
digging existing stems and rhizomes from the soil.
Germination
Seeding rate
Seeds/pound
Planting depth
Seed maturity
15-96%
31,730
0.5”
May-Nov.
* Germination improved to 96% following seed coat scarification
Prairie acacia
Rio Grand Clammyweed (Polanisia dodecandra ssp. riograndensis)
Clammyweed is an annual forb found throughout Southwestern Texas. Clammyweed has bright pink
flowers tha attract a wide variety of butterflies, bees, and other pollinators. Seed from clammyweed
is frequently consumed by game and non-game birds and mammals. The leaves have a rank odor
when crushed.
Germination
50-70%
Seeding rate Seeds/pound Planting depth Seed maturity
March-Oct.
Rio Grand clammyweed
Seacoast bluestem (Schizachyrium scoparium var. littorale)
Seacoast bluestem is the dominant, climax grass species over most of the upland grasslands of
the South Texas Sand Sheet. It is also common on barrier islands along the Gulf Coast. Its range
is restricted to sand or sandy loam textured soils. It provides good to excellent forage for livestock,
and decreases in abundance following continuous grazing. Because of its bunchgrass growth form,
it provides good nesting cover for ground nesting birds, such as bobwhite quail. It grows in large,
dense, colonial stands by means of rhizomes. Two primary growing seasons of seacoast bluestem
occur in early spring and late summer, however green leaves are present throughout much of
the year depending on moisture and temperature. Seedheads initiate growth from July through
September depending on rainfall and reach maturity from October through December. Seed can
be harvested from seacoast bluestem by use of a flail-vac, pull type seed stripper or modified
combine. Planting of seacoast bluestem should be done in late fall or early spring. When planted
using a seed drill, seed should be mixed with some type of carrier to enable uniform planting
and ease of flow through seed tubes and boxes, because awns and small hairs commonly cause
blockages. Seeds can also be broadcasted on a prepared seedbed and covered with a roller or
packer. Seacoast bluestem can also be restored using native hay. Handling of seacoast bluestem
hay should be kept to a minimum to prevent excessive seed shatter.
Germination
Seeding rate
Seeds/pound
Planting depth
Seed maturity
8-30%
7-9 lbs
PLS/ac.
839,475
0.5”
Oct.-Dec.
Seacoast bluestem
Shortspike windmillgrass (Chloris subdolichostachya)
Shortspike windmillgrass is a warm season perrenial grass found throughout South Texas. It is a
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naturally occurring hybrid between Chloris cucullata and Chloris verticillata. This grass is found
in a variety of habitat types. Common use for conservation plantings include grassed waterways,
streamside buffers, filter strips, and pond embankments. Shortspike has shown promise in being
able to establish in stands of buffelgrass, and competes well with it. “Welder” is a South Texas
release of this species available for planting.
Germination
Seeding rate
Seeds/pound
Planting depth
Seed maturity
>90%
0.25-0.50 lbs
PLS/ac
3,000,000
0.12-0.25”
May-Oct.
Shortspike windmillgrass
Sideoats grama (Bouteloua curtipendula)
Sideoats grama is a common, perrenial native grass found in South Texas. It is designated as
the State Grass of Texas. Sideoats is found on gravelly hills, loamy, clayey and sandy loam soils.
Forage value is good, and excellent nesting cover for ground nesting birds is also provided by this
grass.
Germination
Seeding rate
Seeds/pound
Planting depth
Seed maturity
9%
5-6 lbs PLS/ac
168,000
0.25”
March-Oct.
Sideoats grama
Silver bluestem (Bothriochloa laguroides var. torreyana & B. longipaniculata)
Silver bluestem is a very common perennial bunchgrass that is widely distributed throughout South
Texas. Silver bluestem is a fair to good forage for livestock and provides some value as cover for
wildlife. It is a resilient grass that is often times the only native species that persists (in any quantity)
in areas heavily invaded by exotic grasses. Silver bluestem produces seed several times a year
under favorable conditions. It can be easily harvested by hand or mechanically.
Germination
Seeding rate
Seeds/pound
Planting depth
Seed maturity
70-90%
2 lbs PLS/ac
483,600
1/4”
May-Nov.
Silver bluestem
Slender grama (Bouteloua repens)
Slender grama is an early successional native grass found throughout South Texas. Slender
grama has shown excellent competitive ability with many non-native grass species in experimental
plantings. Best uses for revegetation projects include erosion control and highway right-of-way
plantings. “Dilley” is a South Texas release of this species available for planting.
Germination
Seeding rate
Seeds/pound
Planting depth
Seed maturity
33%
8 lbs PLS/ac
116,300
0.25”
March-Oct.
Slender grama
Slim tridens (Tridens muticus var. muticus)
Slim tridens is a warm season, perennial grass found on various sites in South Texas. It is a fair
forage for livestock and produces seed eaten by birds and small mammals. Slim tridens produces
136

seed throughout the year following rainfall.
Germination
Seeding rate
Seeds/pound
Planting depth
Seed maturity
78%
2 lbs PLS/ac
722,528
1/4”
May-Nov.
Slim tridens
Spiny hackberry (Celtis pallida)
Spiny hackberry (or granjeno) is a thorny, evergreen shrub common to most of South Texas. Spiny
hackberry is found in a variety of soils and habitat types. It is an excellent plant for wildlife, and
a variety of mammals and birds consume the seeds. It also provides nesting and loafing habitat
for a variety of birds. The leaves and new shoots are eaten by livestock in periods of drought
when other forage is unavailable. The seeds of spiny hackberry should be cleaned from the fruit
after collection if planting will be delayed. Care should be taken when drying cleaned seeds, as
this species is especially susceptible to molding. Seeds that will be planted soon (1-3 days) after
collection may be planted with the fruit (pulp) intact, with favorable germination. Cold scarification
is also recommended for increasing germination of spiny hackberry. Germination as high as 62%
has been obtained using a combination of mechanical scarification, gibberellic acid and heat/chill
treatments. The plant can also be propagated vegetatively from softwood cuttings.
Germination
≤ 62%
Seeding rate Seeds/pound Planting depth Seed maturity
June-Nov.
Spiny hackberry
Switchgrass (Panicum virgatum)
Switchgrass is a rhizomatous, perennial bunchgrass native to parts of South Texas. It is considered
a climax co-dominant of lowland clay sites and of upland sandy range sites. It is found in two
distinct growth forms. The first being a colonial form with numerous stalks and bright green leaves,
it is found almost entirely on sand or sandy loam soils in the sand sheet. The second is an upright
bushy form found in lowlands along creeks, rivers, and other riparian areas. This form is more
blue-green in color and is restricted to heavy clay and clay-loam soils. Switchgrass is good forage
for livestock and wildlife readily eat the seeds and use the foliage for nesting and loafing cover.
Switchgrass responds well to prescribed fire. It produces numerous seed stalks in late summer
and seed matures throughout the fall. Seed is occasionally produced in spring or early summer if
adequate moisture is available. Production plots of switchgrass produce an average of 300 pounds
per acre.
Germination
Seeding rate
Seeds/pound
Planting depth
Seed maturity
9-67%
2 lbs PLS/ac.
426,000
≤ 0.25”
Sept.-Nov.
Switchgrass
Texas grama (Bouteloua rigidiseta)
Texas grama is a warm season perennial grass common throughout South Texas. It is found in a
variety of soil types and plant communities, commonly on roadsides, overgrazed pastures or other
disturbed sites. Texas grama is a low growing, early successsional plant and has value as a fast
establishing cover on disturbed sites. Texas grama seed matures several times per year, usually
137

following rainfall events.
Germination
Seeding rate
Seeds/pound
Planting depth
Seed maturity
10-95%
9 lbs PLS/ac
103,680
1/4”
April-Nov.
Texas grama
Texas kidneywood (Eysenhardtia texana)
Texas kidneywood is a warm season, perennial shrub found in South Texas. It grows in Texas and
south into Mexico. In Texas, it can be found in the Trans-Pecos, Edwards Plateau, Southern Coastal
Prairie and Rio Grande Plains regions. In most cases, it prefers calcareous soils and is found as
part of brushy, chaparral vegetation. Texas kidneywood prefers full sun or light shade. It is drought
tolerant, but may temporarily defoliate under extended drought conditions. It grows rapidly under
moist conditions. The leaves are browsed by livestock and white-tailed deer. Texas kidneywood
can be grown from seed or from cuttings. Texas kidneywood germinates best when temperatures
fall between 68 – 86ºF and when there is about 12 hours of daylight. Colder temperatures have
been known to reduce germination, whereas higher temperatures tend to reduce the survival
of new seedlings. However, Texas kidneywood has been known to germinate with no light and
temperatures from 15º to 40ºC. Young plants may do better in light shade until they become better
established. Texas kidneywood can also be grown from soft or semi-hardwood cuttings. Cuttings
4 to 6” long should be taken in the summer and early fall. A rooting hormone should be used to
facilitate root growth. Cuttings tend to root in 3 to 4 weeks. Seedpods should be harvested when
they have turned brown and dry. Seeds are kidney-shaped, slightly-plump, and light to darker
brown when matured. Seeds or pods should be dried at room temperature for several days before
storing. It is recommended that seeds or pods be fumigated and stored at cold temperatures.
Germination Seeding rate Seeds/pound
58,520
Planting depth
0.25-0.50”
Seed maturity
June-Oct.
Texas kidneywood
Texas persimmon (Diospyros texana)
Texas persimmon is a shrub or small tree found in a variety of sites in South Texas. It produces
a large black fruit containing several seeds during mid-summer. The fruit is eaten by a variety of
wildlife, deer also browse the leaves. Fruits turn a deep black color when ripe. The seeds should
be cleaned from the fruit upon collection. Freshly collected seeds will readily germinate. Seeds
stored after collected should be cold scarified to break dormancy. Some research suggests that
germination of seeds stored at room temperature is not decreased.
Germination Seeding rate Seeds/pound Planting depth Seed maturity
June-Aug.
Texas persimmon
Texasgrass (Vaseyochloa multinervosa)
Texasgrass is a warm season perennial grass found on sandy sites throughout the South Texas
Sand Sheet. Texasgrass can be found growing in open prairies, live oak mottes and on the edges
138

of other brush canopies. It is endemic to South Texas, meaning it grows nowhere else in the world.
Texasgrass provides good forage for livestock and good cover for wildlife. Seeds from Texasgrass
are readily consumed by bobwhite quail. Texasgrass usually initiates growth in summer and
produces seed in early fall. Dependant on rainfall, seed can also be produced in mid summer.
Texasgrass produces ample seed that can be easily collected by hand or with mechanical seed
strippers.
Germination
Seeding rate
Seeds/pound
Planting depth
Seed maturity
8-26%
174,966
≤ 0.25”
July-Oct.
Texasgrass
Yellow indiangrass (Sorghastrum nutans)
Yellow indiangrass is a warm season, colonial bunchgrass obtaining a height of up to 7’ in South
Texas. It is found along the South Texas Coastal Prairie and Sand Sheet. Other populations exist
along streambeds and drainages in north eastern South Texas. It provides excellent forage for
livestock, and has value to wildlife as cover and nesting habitat. Yellow indiangrass can be found
in solid stands where it is protected from grazing. These dense stands are void of most other
vegetation. Yellow indiangrass spreads vegetatively by means of underground rhizomes and by
seed. The seeds are easily harvested upon maturity because of the long seed stalks which hold
the seed above the foliage.
Germination
Seeding rate
Seeds/pound
Planting depth
Seed maturity
2-15%
5-8 lbs PLS/ac.
129,390
1/2”
Oct.-Nov.
Yellow indiangrass
139

REFERENCES
Correll, D. S., and Johnston, M.C., (1996). Manual of the Vascular Plants of Texas. Richardson, TX: The University of
Texas at Dallas.
Everitt, J.H. (1984). Germination of Texas persimmon seed. J. Range Management. 36:189-191.
Everitt, J.H. and Drawe, D. L. (1993). Trees,Shrubs, and Cacti of South Texas. Lubbock, TX: Texas Tech University
Press.
Everitt, J.H., Lonard, R. I. and Drawe, D. L. (1997). Field Guide to the Broad-leaved Herbaceous Plants of South Texas.
Lubbock, TX: Texas Tech University Press.
Fulbright, T.E., Flenniken K.S., and Waggerman G.L. (1986). Enhancing germination by spiny hackberry seeds. J.
Range Management. 39:552-554.
Hatch, S. L., Gandhi, K.N., and Brown, L.E. (2001). Checklist of the Vascular Plants of Texas. College Station, TX: Texas
Agricultural Experiment Station.
Hatch, S. L., Schuster, J. L., and Drawe, D. L. (1999). Grasses of the Gulf Prairies and Mashes. College Station, TX:
Texas A&M University Press.
Jones, F. B., (1982). Flora of the Texas Coastal Bend. Sinton, TX: Welder Wildlife Foundation.
Kika de la Garza PMC (1999). A Germination Study of Eighteen Accessions of Texas Kidneywood. Technical Note v.2
(6). Kingsville, TX: USDA/NRCS.
Kika de la Garza PMC (2002). 2002 Annual Report. Kingsville, TX: USDA/NRCS.
Kika de la Garza PMC (2003). 2003 Annual Report. Kingsville, TX: USDA/NRCS.
Nokes, J. (1986). How to Grow Native Plants of Texas and the Southwest.
Houston, TX: Gulf Publishing.
South Texas Natives. (2004). Database of STN Seed Collections. Kingsville, TX.
Speer, E.R., and Wright, H.A. (1981). Germination Requirements of Lotebush (Ziziphus obtusifolia var. obtusifolia). J.
Range Management 34:365-368.
Taylor, R. B., Rutledge, J., and Herrera, J. G. (1997). A Field Guide to Common South Texas Shrubs. Austin, TX : Texas
Parks and Wildlife.
Texas Parks and Wildlife Department (2004). Plant Information Database.
http://tpid.tpwd.state.tx.us/species_commonname.asp.
Vora, R.S. (1989). Seed germination characteristics of selected native plants of the lower Rio Grande Valley, Texas. J.
Range Management. 42:36-42.
Whisenant, S.G., and Ueckert, D. N. (1982).
Germination responses of Eysenhardtia texana and Leucaena retusa. Journal of Range Management, 35 (6), 748-750
140

APPENDIX D
Common and Scientific Names
Mentioned in Text
Plants
Authority for scientific names is Hatch S.L., Gandhi K.N., and Brown L.E. 2001 (A Checklist of the Vascular
Plants of Texas. http://www.csdl.tamu.edu/FLORA/taes/tracy/regeco.html)
________________________________________________________________
Common name
Scientific name
Anaqua
Ehretia anaqua
Arizona cottontop
Digitaria californica
Awnless bush sunflower
Simsia calva
Big bluestem
Andropogon gerardii
Blackbrush acacia
Acacia rigidula
Black hickory
Carya texana
Bristle leaf dogweed
Thymophylla tenuiloba
Buffalograss
Buchloe dactyloides
Bulrush
Scirpus spp.
Bundleflower
Desmanthus spp.
Carolina wolfberry
Lycium carolinium
Canada wildrye
Elymus canadensis
Cattail
Typa spp.
Cedar elm
Ulmus crassifolia
Ceniza
Leucophyllum frutescens
Cereal rye
Secale cereale
Clover
Trifolium spp.
Clubhead cutgrass
Leersia hexandra
Coma
Bumelia celestrina
Common curlymesquite
Hilaria belangeri
Common sunflower
Helianthus annuus
Coontail
Ceratophyllum spp.
Cowpea
Vigna spp.
Creosotebush
Larrea tridentata
Desert yaupon
Schaefferia cuneifolia
Douglas fir
Pseudotsuga menziesii
Drummonds phlox
Phlox drummondii
Drummonds skullcap
Scutellaria drummondii
Eastern cottonwood
Populus deltoids
Eastern gamagrass
Tripsacum dactyloides
Engelmann daisy
Engelmannia pinnatifida
Fall witchgrass
Digitaria cognata
False rhodesgrass
Chloris crinita
Firs
Abies spp.
Forage sorghum
Sorghum spp.
Fringeleaf paspalum
Paspalum setaceum var. ciliatifolium
Gayfeather
Liatris punctata
Granjeno
Celtis paillada
Guajillo
Acacia berlandieri
141

Gulf cordgrass
Gulfdune paspalum
Guayacan
Mesquite (Honey mesquite)
Hairy grama
Hogplum
Hooded windmillgrass
Huisache acacia
Indian blanket
Inland seaoats
Juniper
King Ranch bluestem
Kleberg bluestem
Little bluestem
Littleleaf sumac (evergreen sumac)
Live oak
Longtom paspalum
Lotus
Marshhay cordgrass
Medic
Millet
Mormon tea
Mountain laurel
Multiflowered false rhodesgrass
Muscadine
Muskgrass
Mustang grape
Oats
Partridge pea
Pink pappusgrass
Plains bristlegrass
Prarie acacia
Pecan
Pine
Post oak
Sago palmweed
Seacoast bluestem
Seashore dropseed
Seaoats
Sedge
Scotch thistle
Shagbark hickory
Shortspike windmillgrass
Shrubby oxalis
Sideoats grama
Silver bluestem
Smooth cordgrass
Sorghum
Soybean
Spanish dagger
Spiny aster
Spruce
Sudangrass
Spartina spartinae
Paspalum monostachyum
Guaiacum angustifolium
Prosopis glandulosa
Bouteloua hirsuta
Colubrina texensis
Chloris cucullata
Acacia smallii
Gaillardia pulchella
Chasmanthium latifolium
Juniperus spp.
Bothriochloa ischaemum
Dichanthium annulatum
Schizachyrium scoparium
Rhus microphylla
Quercus virginiana
Paspalum lividum
Nelumbo lutea
Spartina patens
Medicago
Setaria spp.
Ephedra antisyphilitica
Sophora secundiflora
Chloris pluriflora
Vitus rotundifolia
Chara spp.
Vitus mustangensis
Avena sativa
Chamaecrista fasciculata
Pappophorum bicolor
Setaria leucopila
Acacia angustissima
Carya illinoinensis
Pinus spp.
Quercus stellata
Potamogeton pectinatus
Schizachyrium scoparium var. littoralis
Sporobolus virginicus
Uniola paniculata
Carex spp.
Onopordum acanthium
Carya ovata
Chloris x subdolistachya
Oxalis berlandieri
Bouteloua curtipendula
Bothriochloa laguroides
Spartina alterniflora
Sorghum spp.
Glycine max
Yucca treculeana
Leucosyris spinosa
Picea spp.
Sorghum bicolor
142

Sugar hackberry
Sunflowers
Switchgrass
Tanglehead
Texas ebony
Texas mountain laurel
Texas persimmon
Texas prickly pear cactus
Texas varilla
Texas wintergrass
Tricale
Twisted acacia
Vetches
Vine ephedra
Water nymph
Water stargrass
Wheat
Whiplash pappusgrass
Whitebrush
Wigeongrass
Wild onion
Winter peas
Wooly globe mallow
Celtis laevigata
Helianthus spp.
Panicum virgatum
Heteropogon contortus
Pithecellobium flexicaule
Sophora secundiflora
Diospyros texana
Opuntia lindheimeri
Varilla texana
Stipa leucotricha
x Triticosecale
Acacia schffneri
Vicia spp.
Ephedra antisyphylitica
Najas spp.
Heteranthera dubia
Triticum spp.
Pappophorum vaginatum
Aloysia gratissima
Ruppa maritima
Allium drummondii
Pisum spp.
Sphaeralcea lindheimeri
Mammals
Common and scientific names of mammals mentioned in text. Authority for scientific names is: Davis, W.B.
1960 (The Mammals of Texas, Texas Game and Fish Commission, Austin, Texas).
________________________________________________________________
Common name
Scientific name
Ocelot
Felis pardalis
White-tailed deer
Odocoileus virginianus
Birds
Common and scientific names of birds mentioned in text. Authority for scientific names is: American
Ornithologists Union 2004 (AOU Checklist of North American Birds, http://www.aou.org/aou/birdlist.html).
Common name
Northern bobwhite quail
Northern shoveler
Piping plover
Sandhill crane
Scientific name
Colinus virginianus
Anas clypeata
Charadrius melodus
Grus canadensis
143

APPENDIX E
Construction of Shrub Nursery
Developing your own nursery for the propagation of native plants is a feasible plan, even if you
are not a commercial grower. There are several steps involved with this process. The design of
the nursery presented here is intended to be practical, with an effort to reproduce natural growing
conditions. Seedlings that will be transplanted into the field within the same year will need to
survive in considerably less than ideal conditions. Therefore,
the nursery is open-air, with individual shade-cloth support
structures for each bench. Furthermore, mild winters and
unbearably hot summers in South Texas somewhat eliminate
the need for enclosed greenhouses.
Nursery Location
The following factors should be considered when choosing a
nursery location:
• Water availability
• Electricity sources (if necessary)
• Amount of sunlight
• Aspect (orientation to the path of the sun)
• Access both by foot and with vehicles
• Convenience to storage areas and shaded or enclosed
workspaces
• Eventual capacity (size)
Shrub nursery
The nursery should be located in full sun, so that the amount of sunlight reaching the seedlings
can be controlled. Any shade from adjacent structures or trees during any time of the day may
complicate the control of lighting, watering schedules, fungus growth and other diseases, and the
general health of the seedlings.
Nursery Layout
A well thought-out layout for the nursery should include the location and dimensions of individual
benches, water lines, and electrical lines (if desired). The most efficient design will place benches
in short rows of 2 to 4 benches with wide aisles between rows. If you have sufficient space, leave
room at the end of each or every second bench as a pass-through, so that workers don’t have to
walk all the way to the end of a row in order to get to the other side. When deciding on the aisle
widths, consider the width of any equipment (wheelbarrow, nursery cart, ATV) which you may be
using in the greenhouse.
144
Ground Preparation
The ground on which the nursery is constructed should be free
of all vegetation (bare dirt), as level as possible, and slightly
soft at the time of construction for leveling purposes (Photo).
Any underground plumbing or electrical wiring needed for the
nursery should be installed first and tested for leaks; trenches
should be backfilled and firmly packed and leveled. Standard
nursery ground cloth can then be applied to the entire area
Leveled ground overlaid with ground cloth

and anchored in place. A good drainage system around the perimeter of the area can be easily
constructed by digging a shallow trench to drain excess water away.
Nursery Bench Design
Once your ground is prepared, your next step will be to set up the nursery benches. These are
portable, inexpensive, and easy to construct so that expansion can take place as needed. The
base structure is designed so that it can be easily dismantled and moved. Each bench consists of
3 main parts:
• A base of 8”x8”x16” hollow core concrete blocks used to form columns which raise the
bench to a comfortable working level
• A platform to hold the plants
• A structure to support the shade cloth and an overhead watering system
The base structure must be strong and stable to support the overall weight of the filled planting
containers, and the platform must be able to support up to 1 ton of soil mix. Platforms must also
be designed to allow for adequate water drainage and air circulation for air-pruning the roots. The
overhead structure must be strong enough to resist average winds, tall enough to allow comfortable
access underneath, removable, and versatile enough to support either shade cloth or plastic
sheeting. All parts must be water and rot-resistant (plastic or treated wood).
Equipment List
Bench base instructions:
Place the first hollow block in each column with the voids facing up, and not to the side. The bottom
block of each column should be placed exactly according to the dimensions shown in Figure 1.
• Each block will need to be perfectly level, independently and in relation to the others.
Since the blocks are placed on ground cloth, leveling can be difficult if the ground
underneath is not slightly soft. Soil can be leveled through the ground cloth using the flat
end of the block itself, or some other leveling tool. This is the most time-consuming and
painstaking part of the construction, but leveling is important to ensure that the bench
does not begin to lean and collapse when it is bearing the full weight of the soil.
• Once the bottom blocks are accurately placed and leveled, 2-3 blocks are then stacked
on top of each bottom block and checked again for leveling.
• To anchor columns, drive 1” diameter galvanized pipe or rebar through the voids in the
stacked blocks, puncturing the ground cloth, and penetrating the ground by at least one
foot. The pipe should ultimately be equal to or below the upper rim of the topmost block
(Fig. 2).
to outside
edge
96” (8’)
to center
16”
(inside
edge to
inside
edge)
to outside
edge
28”
to center
28”
72”
inside edge to inside edge
Figure 1. Placement of concrete blocks for nursery bench case - top view
to outside
edge
145

landscape timbers
ground level
blocks placed with voids in
a vertical orientation
Figure 2. Placement of concrete blocks for nursery bench base - side view
columns are anchored with
1” diameter galvanized
pipe or rebar driven
through the voids in the
blocks and into the ground
to a depth of 1’
Bench platform instructions:
• Place each pair of the 6 8-foot long wood landscape timbers
lengthwise end-to-end, on top of each of the 3 rows of block
columns (Fig. 2).
• A 16’ x 52” stock panel is then placed on top of the landscape
timbers and fastened in place using fence staples.
• A 16’ length of 48” (¼” square) hardware cloth is then placed on
top of the stock panel and wired to the stock panel at each end.
• Construct rectangular, bottomless boxes to hold the planting
containers using (2) 2” x 6” x 4’ treated wood boards for the ends
and (2) 1” x 6” x 8’ boards for the sides. Reinforce the ends with
angle braces. Place each box end-to-end on top of the hardware
cloth (Fig. 3).
Shade Cloth and Support Structure
Shade cloth helps moderate temperatures in the South Texas heat
Figure 3. Bottomless boxes placed
end to end
during the initial growth period between spring-fall. The partial shade also helps to conserve water,
as plants may need to be watered up to 2 times a day in the month of August. A density of 30% for
the overall nursery is sufficient, but as much as 70% shade may be needed for recently transplanted
seedlings. If necessary, simply reserve a bench with a higher density shade cloth. Shade cloth
also helps to induce faster elongation of the main stem of the seedling. Because the plant growth
hormones that “plasticize” the dividing cells in the apical meristem are photo-sensitive, a reduction
in light intensity results in longer cells. If shade is excessive, however, the seedlings will become
spindly and weak-stemmed. Approximately 10-15 days before planting, seedlings should gradually
be exposed to more sunlight to help them adapt and survive in the field. Simply remove the shade
cloth, or reduce the amount of shade.
The overhead cloth support structure is constructed out of Schedule 40 (3/4”) PVC pipe, and
consists of 5 “ribs” that form a tunnel over the greenhouse (Figure 4). Each rib is connected by
horizontal PVC braces that are attached through elbows and crosses along the tops and sides of
the ribs. The pipe is fastened to the outside of the planting boxes. This design also allows for the
146

attachment of water lines and sprinklers
to the underside of the overhead frame, if
desired. The shade cloth is simply draped
over the structure and anchored in place
with temporary fasteners.
For the overhead cloth support structure,
you will need the following supplies:
• Pipe segments (Schedule 40 – ¾”) •
Hardware and Other Supplies
• 20 segments 8” long
• 1 can PVC glue, 4-oz
• 10 segments 12” long
• 1 length 30% shade cloth, 25’ x 12’
• 10 segments 36” long, for legs
• 20 each ¾” 1-hole EMT clamps
• 12 segments 46” long, for horizontals
between ribs
• 20 each 1” long, gasketed (“Neowasher”)
¼”metal-to-wood screws
• 30 lengths baling twine
• 24” PVC fittings
• 6 tees
• 9 crosses (4-way connectors)
• 20 elbows, 45º
8”
8”
leg
Figure 4. Shade frame assembly - diagram of ribs and ends
Overhead cloth support structure
instructions:
• Cut the 10’ lengths of PVC pipe according to the above list.
• Construct each rib, and connect them together using the horizontal sections that fit into the
crosses and elbows.
• Once the frame is standing, and appears to be symmetrical, glue the joints by disassembling and
gluing one joint at a time, making sure to keep each joint at its proper angle.
• Once the frame is complete, center the PVC structure over the benches, with the legs of the
frame on the outside.
• Using 1” rubber-gasketed hex-head (“Neowasher”) screws, attach the EMT clamps across each
PVC frame leg to the side of the planting box. Space one clamp near the bottom of the box, and
the second toward the top. The bottom of the frame leg should be flush with the bottom edge of
the box. (Photos)
• Nail a fence staple (1 ½”) to the center of each box end, and to the center of the box sides for the
benches that are located in between the legs of the shade frame. Nail each fence staple only
halfway because they will be used to secure the twine used for attaching the shade cloth to the
frame.
Shade cloth instructions:
Before attaching the shade cloth to the frame, cut at least 30 pieces of baling twine about 24” long.
Tie the middle of one piece using a simple knot to each fence staple in the sides and ends of the
boxes so that both ends of the twine are free; these will anchor the shade cloth edges when the
8”
end
12” 12”
Ends - 2 exactly alike
12” 12”
8” 8”
Ends - 2 exactly alike
rib
rib
end
rib
assembled frame
8”
tee
cross
8”
8”
horizontals between ribs
45 o
elbow
36” leg
147

sides are rolled down. Tie one piece of twine to the middle of each 46” horizontal segment of PVC
pipe on the frame; tie twine at middle so that both ends are free; these will anchor the shade cloth
when the sides are rolled up. Place the shade cloth over the frame, making sure it is centered.
At the very top of the frame, in 3 or 4 places, thread one end of twine through the cloth, over
and around the PVC and back down through the cloth; secure the twine using a bowknot on the
underside of the PVC to fasten the shade cloth securely to the frame. Roll up the sides and ends
of cloth to the first horizontal PVC segments; thread one end of twine through and secure with a
bowknot; this will be the position of the shade cloth when you are working with the plants.
Watering System
Overhead systems can be economically and easily installed for automatic watering of the nursery.
Temporary systems can last over 5 years, but more permanent systems can be installed with the
help of a nursery supply house or a professional irrigation system installer. In all cases, ensure that
the water supply is accessible to each bench.
Important Notes:
▪ Pipes running from the main water line should be smaller than that of the main line to increase
the water pressure as it approaches the tables.
▪ Tables should be set up as close to the main water line as possible to allow for maximum water
pressure.
Instructions:
▪ Develop a well-planned and efficient design; water lines will tee-off from a main water line, and
then emerge from the ground at the end of each bench.
▪ Use a 2” PVC pipe for the main supply, but reduce the size to 1” for individual water lines leading
to each bench.
▪ Prepare the trenches and lay out the water lines, but do not glue until the lines are laid out and
checked for size and fitting.
▪ Using a 90 o elbow, create a vertical extension of approximately 2’ at the end of each bench.
Attach a faucet in a horizontal position so it is easy to use, and install an automatic timer and
regulator just above the cut-off valve.
Design Option 1:
▪ Materials List (per bench):
▪ Water supply line to bench, installed
▪ Faucet, incl. fitting(s) to install to water line
▪ Watering timer, battery-powered, electronic, programmable, 4-cycle
▪ Pressure regulator
▪ Female header-to-supply connector
▪ Header hose, black poly, 1/2” diameter
▪ Multi-pattern sprinkler assembly
▪ End cap
▪ Hole punch with goof plugs
▪ Carefully roll out the poly-header pipe, taking care not to kink the tubing.
▪ Insert the cut end of the header pipe into a double-ended male threaded compression coupling;
screw the end of the cap onto the free end of the coupling. This will be the terminal end of the
148

ench’s sprinkler line.
▪ Using twine or some other fastener, secure the header pipe to the underside of the top of the
shade frame, starting with the terminal end; secure the line in 5 or 6 places.
▪ Allow the supply end of the pipe to curve gently to the side and down along the corner shade
frame support so it can be connected to the water supply, and then fasten in place.
▪ Cut the supply end of the header pipe to length at a slanted angle, insert the end into the
compression end of the female threaded hose end coupling, and then screw the female threaded
coupling onto the regulator.
▪ Using a hole-punch, punch 5 holes in the bottom side of the header pipe above the bench; one
near each end, one in the center, and the other two spaced between the middle and ends.
▪ Insert one end of the connector into a riser tube of a multi-pattern sprinkler and the other end into
one of the holes in header pipe. The sprinkler assemblies should point straight down; adjust if
needed.
▪ Sprinklers will drip for several minutes after each watering, and can sometimes wash out seeds
in the containers directly below. To prevent this, a string can be tied from the sprinkler head
extending to the bench between containers, which will allow the drip to run along the string and
away from the seed containers.
Design Option 2:
Materials List (per bench):
▪ 2” PVC pipe
▪ 1” PVC pipe
▪ ¾” PVC pipe
▪ 2” tees
▪ 1” 90 o elbows
▪ ¾” tees
▪ 2” > 1” reducers
▪ 1” > ¾” reducers
▪ Risers; various lengths depending on plant height
▪ ¾” pipe clamps
▪ Pipe cutters
▪ Trenching shovels
▪ Primer
▪ Pipe glue
▪ Teflon® tape
▪ 1” cut-off valves or faucets
▪ Watering timer, battery-powered, electronic, programmable, 4-cycle
▪ Adjustable sprinkler heads with minimum 6’ range
▪ Install a “tee” above the timer/regulator, and run ¾” PVC pipe around the perimeter of the table;
do not connect, but cap the ends or install a 90 o elbow if a sprinkler will be attached. Attach to
wood frame with clamps if necessary.
▪ Attach “tees” where sprinkler heads will be placed, and fit with reducers for ½” PVC pipe.
▪ Attach ½” risers, and install sprinkler heads.
▪ After pipes have been glued, test the system for leaks. The trenches can then be buried, and
the timers set for the appropriate watering schedule. Sprinkler patterns and plant vigor should
be checked regularly.
149

APPENDIX F
Resources
CONSULTANTS
USDA-NRCS
Rangeland Management
Specialists see:
http://www.nrcs.usda.gov
Carter Ranch Consulting
Cliff Carter
234 Lakeview Drive
Victoria, TX 77905
Ph: 361-578-9296
E-mail: cwcarter@zamigo.net
Environmental Survey. Inc.
David Mahler
4602 Placid Place
Austin, TX 78731
Ph: 512-458-8531
Fax: 512-458-1929
E-mail: dvmahler@envirosurvey.com
Web site: www.envirosurvey.com
Paula Maywald
PO Box 16
Kingsville, TX 78364
Ph: 361-522-8128
Neiman Environments, Inc.
Bill Neiman, Jay Kane
Mail Order Station 127 N 16 th St.
Junction, Texas 76849
Ph: 800-728-4043
Fax: 325-446-8884
E-mail: info@seedsource.com
Web site: www.seedsource.com
Stan Reinke
203 Madera
Victoria, TX 77905
Ph: 361-570-0228
Resource and Land Management, Inc.
Mike Peters
204 Brian Drive
Pleasanton, TX 78064
Ph: 830-569-6826
Fax: 830-281-8188
E-mail: mapag@texas.net
Whisenant Ecological Services
Dr. Steve Whisenant
15122 Post Oak Bend
College Station, TX 77845
CONTRACT GROWERS & LANDSCAPE
SUPPLIERS
Speedling Inc.
Alamo Transplant Nursery Division
P. O. Box 730
Alamo, TX 78516
Ph: 956-787-1911
Ph: 800-892-5266
Fax: 956-787-5556
E-mail: Texas@speedling.com
Web site: www.speeding.com
Mike Heep
1714 S. Palm Court Dr.
Harlingen, TX 78552
Ph: 956-381-8813
E-mail: heep03@yahoo.com
Lomitas Nursery
Benito Trevino
P.O. Box 422
Rio Grande City, TX 78528
Ph: 956-486-2576
E-mail: lomitas@vsta.com
Madrone Nursery
Dan Hosage
2318 Hilliard
San Marcos, TX 78666
Ph: 512-353-3944
E-mail: madronenursery@earthlink.net
McNeal Growers
Pat McNeal
P.O. Box 371
Manchaca, TX 78652
Ph: 512-280-2233
Fax: 512-291-5220
E-mail: information@mcnealgrowers.com
Web site: www.mcnealgrowers.com
150

NATIVE SEED DEALERS
Bamert Seed Company
1897 CR 1018
Muleshoe, TX 79347
Ph: 806-272-5506
E-mail: nbamert@bamertseed.com
Web site: www.bamertseed.com
Douglass W. King Co.
Dean Williams
P. O. Box 200320
San Antonio, TX 78220-0320
Ph: 210-661-4191
Ph: 1-888-DK-SEEDS
Fax: 210-661-8972
E-mail: sales@dkseeds.com
Web site: www.dkseeds.com
Native American Seed
Bill Neiman, Jay Kane
Mail Order Station 127 N 16 th St.
Junction, Texas 76849
Ph: 800-728-4043
Fax: 800-728-3943
E-mail: info@seedsource.com
Web site: www.seedsource.com
Pogue Agri Partners
Keith Walters & Gary Pogue
P. O. Drawer 389
Kenedy, TX 78119
Ph: 830-583-3456
Fax: 830-583-9843
E-mail: pogueagri@aol.com
Web site: www.pogueargri.com
Turner Seed Company
Darcy Turner, J Mercer
211 CR 151
Breckenridge, TX 76424-8165
Ph: 254-559-2065
Ph: 800-722-8616
Fax: 254-559-5024
E-mail: darcy@texasisp.com
Web site: www.turnerseed.com
Watley Seed Company
Andy Watley
Box 51
Spearman, TX 79081
Ph: 806-659-3152
Ph: 1-800-338-2885
E-mail: watleyseed@valornet.com
TURF PRODUCERS
Bladerunner Farms Inc.
David Doguet
802 Howard Rd.
Poteet, TX 78065
Ph: 830-276-4455
Ph: 888-717-4455
Fax: 830-276-8618
E-mail: info@bladerunnerfarms.com
Web site: www.bladerunnerfarms.com
SEED COATING
Agricoat Seed Services
3021 W. Dakota, Suite 109
Fresno, CA 93722
E-mail: agricoatss@yahoo.com
Web site: www.agricoat.com
Seed Biotics
818 Paynter Ave.
Caldwell, ID 83605 USA
Ph: 208-455-0578
Web site: www.seedbiotics.net
Seed Dynamics
P.O. Box 6069
Salinas, CA 93912 USA
Ph: 831-424-1177
Web site: www.seeddynamics.com
LABORATORIES FOR SOIL, FORAGE &
COMPOST TESTING
A& L Plains Agricultural Laboratories, Inc.
P. O. Box 1590
Lubbock, TX 79408
Ph: 806-763-4278
Fax: 806-763-2762
Web site: www.al-labs-plains.com
BBC Laboratories, Inc.
1217 North Stadem Dr.
Tempe, AZ 85281
Ph: 480-967-5931
Fax: 480-967-5036
E-mail: info@bbclabs.com
Web site: www.bbclabs.com
151

Soil Foodweb, Inc.
Elaine Ingham
1128 NE 2 nd Street, Suite 120
Corvallis, OR 97330
Ph: 541-752-5066
Fax: 541-752-5142
E-mail: info@soilfoodweb.com
Web site: www.soilfoodweb.com
Texas A&M University
Soil, Water and Forage Testing Laboratory
2474 TAMU
Room 345 Heep Center
College Station, TX 77843-2474
Ph: 979-845-4816
Fax: 979-845-5958
MICROBIAL INOCULUMS
Texas Environmental Providers, Inc.
Jim Moore
P. O. Box 42116
Houston, TX 77242-2116
Ph: 832-252-1921
Fax: 832-251-7668
E-mail: jadomo@houston.rr.com
Texas Plant & Soil Lab, Inc.
Esper Chandler, Noel Garcia
5115 W. Monte Cristo
Edinburg, TX 78539
Ph: 956-383-0739
Fax: 956-383-0730
E-mail: ngarcia@tpsl.biz
Web site: www.tpsl.biz
Laboratorios A-L de Mexico, S.A. de C.V.
Ricardo Michel
Esmeralda # 2847 Verde Valle
Guadalajara, Jalisco, Mexico 44550
Ph: (3) 121 7925
333 122 8413
Fax: 333 647 3269
E-mail: lab_al@mexis.com
Web site: www.al_labs.com.mx
152

APPENDIX G
South Texas Natives
Editorial Review
• Fred Bryant, Ph.D., Caesar Kleberg Wildlife Research Institute, Texas A&M
University-Kingsville
• Lynn Drawe, Ph.D., Rob & Bessie Welder Wildlife Foundation
• Barry Dunn, Ph.D., King Ranch Institute for Ranch Management, Texas A&M
University-Kingsville
• Timothy E. Fulbright, Ph.D., Caesar Kleberg Wildlife Research Institute, Texas A&M University-
Kingsville
• C. Wayne Hanselka, Ph.D., Texas Cooperative Extension, Department of Rangeland Ecology
and Management, Texas A&M University System
• David Mahler, Environmental Survey, Inc.
• John Lloyd-Reilley, USDA/NRCS E. Kika de la Garza Plant Materials Center
• Stan Reinke, USDA/NRCS, retired
• Steve Windhager, Lady Bird Johnson Wildflower Center
Editorial Assistants
• David Graves
• Marc Hall, USDA Wildlife Services
• Mari-Vaughan Johnson, Ph.D., USDA-ARS Grassland, Soil and Water Research Laboratory
• Shelly Maher, USDA-NRCS E. Kika de la Garza Plant Materials Center
• C. Michelle Peters
South Texas Natives Co-Chairs
• Katharine Armstrong-Love, Katherine Armstrong, Inc.
• Will Harte, Cerrito Prieto Ranch
153