Native Seed Production Manual

Native Seed Production Manual parkecag Tue, 12/19/2023 - 11:24

About the Manual

The intent of this manual, once completed, is to provide basic information for native seed production of nearly 100 species of the tallgrass prairie flora of the upper Midwest. The information presented is compiled from published accounts coupled with native seed production experience at the Tallgrass Prairie Center at the University of Northern Iowa. Critical to this effort were publications from the USDA-NRCS Plant Materials Program, published research articles and technical notes, personal communication with other native seed growers, and Internet resources. Botanical nomenclature follows Flora of North America, wherever possible. Common names follow USDA Plants Database.

The first edition of this manual was written in 2007 with support from USDA NRCS, the Tallgrass Prairie Center, University of Northern Iowa, and the Iowa Crop Improvement Association. A pdf of the original manual can be downloaded here.

The updated and expanded second edition is being produced as an eBook and made available for free on the Tallgrass Prairie Center website. For those who prefer a print copy, printable versions (pdf) of each general information chapter, species production guide, and appendix table will be provided. You are welcome to download and print the pdf versions to create a printed manual that includes the material of greatest importance to you.

This manual is a work in progress, and new information will be added as it is completed.

Funded by:

Iowa Living Roadway Trust Fund

University of Northern Iowa

Written and Edited by:

First edition (2007)

Greg A. Houseal, Program Manager, Iowa Ecotype Project

‌PDF Native Seed Production Manual (2007)

Second edition (2024)

Revised and expanded by Laura Fischer Walter, Plant Materials Program Manager

in collaboration with Green Iowa AmeriCorps members Bri Hull, Mallory Sage and Andrea Fager

and UNI students Laura Spies, Josie Hutchings and Alex Hanson

General Information

General Information parkecag Tue, 12/19/2023 - 11:30

The first part of the manual consists of four chapters providing general information on greenhouse propagation, harvesting, seed processing, and issues of seed source and quality. Specific examples referencing native species are given to help illustrate the application of the techniques being described.

Propagation of Native Species

Propagation of Native Species parkecag Wed, 01/17/2024 - 09:38

Refer to Appendix A Table 1A [to be added soon] for establishment recommendations for individual species.

Propagation of native species through agronomic methods is necessary to scale up seed supplies for restoring native vegetation in highly fragmented landscapes such as the Midwestern Corn Belt of the US. However, information and techniques for propagating the diverse species needed for restoration are often in short supply, or are closely held secrets of private native seed producers. Here we present methods gleaned from a variety of sources, many of which we have used at the Tallgrass Prairie Center in production of source-identified stock seed for regional native seed growers.

This chapter focuses on establishment of production plots using transplants grown in a greenhouse. This approach has advantages: making efficient use of small amounts of stock seed and achieving more reliable seedling establishment (hence, efficient use of prepared plot space). However, it requires specialized facilities and involves considerable labor. Direct seeding is an effective method for establishing some native prairie species, notably the dominant grasses. Preparing a clean, weed-free seed bed and ensuring that seed is planted at the right depth (i.e., not buried too deeply) are crucial. In the species production guides, we provide estimates of the amount of seed needed, row spacing, and planting depth for direct seeding. However, we do not have experience with direct seeding all of the species and include a disclaimer to that effect when applicable.

Most of the species produced at the TPC have been grown in single-species plots of up to ½ acre or in 4-ft wide rows up to 150-ft long. Some growers produce seed using native or introduced grasses as cover crops within forb rows, in mixed cropping systems of a few species, or by reconstructing prairies for the purpose of seed harvest.

Wild species grown for seed production may show signs of adaptation to production conditions over several generations. At each step in propagation, there is potential for selection for genetic traits that improve fitness in an agronomic production system but may alter the ability of the seed to produce plants capable of establishing, surviving, and reproducing in the wild. Within each of the following sections, we will point out the risk of unintended selection and strategies for avoiding it. These strategies are also summarized in the Chapter: Seed Source and Quality. They will sometimes be in tension with the need for private seed companies to produce a profitable crop.

Seed Dormancy

Dormancy describes seed that does not germinate even when presented with favorable conditions of moisture, temperature, and light. Dormancy is an adaptive trait, allowing germination to occur over time and in the proper season. This vital trait prevents the germination of all seeds at a time that might be suboptimal, or even lethal, for seedling establishment. Staggered germination over time is normal, even with stratification, and should be expected when propagating native prairie species.

The benefits of removing dormancy are twofold: First, more seeds germinate in a shorter period of time, which means limited and costly greenhouse space is used more efficiently. Second, increased germination means that more individual plants will potentially establish, flower, and reproduce, contributing their genetic diversity to the next generation.

There are two main categories of dormancy – primary and secondary. Primary dormancy occurs when seed is dormant upon dispersal, which is typical of many prairie species. Species with secondary dormancy produce seed with the ability to germinate readily upon dispersal (fresh); however, the seed may enter a dormant state if conditions aren’t favorable. Many woodland spring ephemeral species belong to this category of dormancy.

Within the two primary categories, there are several types of seed dormancy, and appropriate strategies are required to remove each type (Table 1). The most common type of dormancy in prairie plants of the grass and sunflower families is physiological dormancy, resulting from biochemical compounds that inhibit germination. The compounds may be produced in the seed itself or translocated to the seed from the plant prior to dispersal. Abscisic acid (ABA), for example, prevents premature germination of mature seeds in the seed head, before dispersal from the parent plant. Concentrations of germination inhibitors, like ABA, decline over time, allowing the seed to germinate. Physiological dormancy may respond to chemical treatments such as gibberellic acid or ethylene which counteract ABA. Another type, physical dormancy, is due to a physical characteristic of the seed; for example, the seed coat may be hard or waxy or otherwise impermeable to water and gas exchange, thus inhibiting germination. Species in the sumac, legume, geranium, and buckthorn families have these characteristics. Seeds with physical dormancy require scarification to remove these barriers. A third type is morphological dormancy: the embryo within the seed is underdeveloped upon dispersal, and warm, moist conditions are generally necessary for maturation (55 to 65 °F, 13 to 18 °C). Species with this type of dormancy are found in the parsley, buttercup, arum, lily, and iris families, among others (Baskin and Baskin 1998). Many seeds may have a combination of morphological and physiological dormancy types, i.e., morphophysiological dormancy, a subtype of which is sometimes referred to as double dormancy. Seeds of these species are often slow to germinate and may require lengthy stratification or repeated stratification at different temperatures. Examples of morphophysiological dormancy are found in the lily, gentian, buttercup, and parsley families. Combinational dormancy means that seeds have both a physical barrier to germination and biochemical inhibitors. Such seeds need both scarification and stratification in order to germinate. Some legumes and members of the borage and buckthorn families fit in this category.

Table 1. Types of dormancy associated with family groups and strategies for breaking dormancy (Adapted from Baskin and Baskin 1998).

Type of Dormancy	Cause	Removed by
Physiological	Biochemical inhibitors of germination (i.e., abscisic acid)	Cold and/or warm stratification, treatment with gibberellic acid (GA) or ethylene, light or dark conditions
	Most common form of dormancy in the grass and sunflower families
Physical	Seed coat impermeable to water/gases	Scarification to simulate natural breakdown of barriers to water uptake
	Found in legume, sumac, geranium, and buckthorn families
Morphological	Underdeveloped embryo	Conditions for embryo development, usually warm, moist conditions
	Found in parsley, buttercup, arum, and lily families
Morphophysiological	Underdeveloped embryo and biochemical inhibitors	Lengthy stratification or a sequence of stratification periods at different temps; some species respond to GA
	Found in gentian, buttercup, parsley, and lily families
Combinational	Physical barriers and biochemical inhibitors	Scarification (including wet heat) and stratification
	Found in borage, buckthorn, and some members of legume family

Choosing Dormancy Breaking Treatments

When planning to propagate native species for seed production, it is important to consider the likelihood of seed dormancy and choose methods to effectively reduce it in that species. Within a plant species and population, individuals vary in their degree of dormancy due to complex genetic traits. If dormancy is not broken, individuals carrying this trait will be eliminated from the production population.

Several sources for finding dormancy breaking information are listed in the references for this chapter, including scientific publications, native plant nursery websites, and a searchable plant propagation database. Information in these sources comes from either scientific studies of germination or practical experience, or both. When different sources disagree significantly, it may be advantageous to divide the seed and subject different samples to each of the suggested treatments. It is also wise to consider the geographic source of the information, as levels of dormancy may differ within a species along a north-south gradient of its natural range (or altitude in other regions). Use caution in applying suggestions from native plant nursery websites: dormancy is a genetic trait that can be diminished in a production population over a few generations unless the grower has applied dormancy-breaking strategies.

For some species, little to no information is available. In those cases, first consider the natural timing of seed dispersal for the species. If its seed typically disperses in fall, cold stratification of different lengths is a good starting point. You may also find protocols for closely related species, divide the seed, and try a range of treatments based on what has worked for other members of the genus. Baskin and Baskin have published a more empirical method called the ‘move-along experiment’ that helps to determine the set of temperature treatments needed to germinate one of these unknowns. If you manage to ‘crack the code’ on a difficult species, consider sharing your techniques on the RNGR Propagation Protocols Database.

Seed Treatments

Scarification

Scarification is a technique that simulates the natural disintegration of the seed coat to initiate germination. A hard or waxy coat will not allow the seed to soak up (imbibe) the water needed for germination until the seed coat breaks down. Seed is scarified either through natural processes such as weathering, abrasion, or partial digestion, or through artificial techniques. Seeds have natural openings for water uptake and these weather or wear away first, especially in seeds with hard seed coats, allowing the seed to imbibe water so germination can occur. The trick of scarification, then, is to accelerate the process of weathering these natural openings so seeds can imbibe water, but stopping short of damaging the seed. Some simple scarification techniques are presented here.

Sandpaper wood blocks – These blocks can be constructed using rubber cement to glue a sheet of fine-grain sandpaper to each of two flat plywood boards. Lay one sand block on a tray and use light pressure and a circular motion to move the other sand block on top of a quantity of seed sandwiched in between the two blocks. Adequate scarification is achieved after a minute or two, when seed begins to look dull.

Percussion scarification – Seeds are shaken vigorously inside a heavy glass bottle for a few minutes. Allow ample room for all of the seeds to impact the sides of the bottle. This technique is considered less aggressive and less likely to damage seeds than the sand block method. In a variation of percussion scarification, a pneumatic paint shaker was modified by Khadduri and Harrington (2002) to scarify the very hard seeds of native locust tree species (Robinia neomexicana, R. pseudoacacia).

Wet heat – Pour boiling water (212°F, 100°C) over the seeds just enough to cover them and allow to cool to room temperature, or immerse seeds in boiling water for five to twenty seconds and remove to rinse and cool. This technique is reserved for certain species and is not broadly recommended. Some species will also require stratification after wet heat. This is effective for New Jersey tea (Ceanothus americanus), hairy puccoon (Lithospermum caroliniense, followed by 90-day cold stratification), and reportedly for false gromwell (Onosmodium molle).

*Image: Hairy puccoon, Lithospermum caroliniense, nutlets subjected to boiling water scarification prior to being cold/moist stratified for 90 days.*

Commercial scarifiers are also available from seed equipment manufacturers, such as a Forsberg scarifier, a sandpaper-lined cylinder with metal paddles that turn and agitate the seed. This is a very aggressive method and only a few seconds are generally needed. Precious seed can be reduced to flour if left on too long. Seed may already have been scarified as part of the cleaning process (e.g., if a scarifier/brush machine was used for dehulling legume seeds). Contact your seed vendor/producer to determine if the seed has been cleaned with a scarifier/brush type machine.

*Image: A Forsberg scarifier, to be used with caution.*

Stratification

Stratification is a process whereby seeds are placed in a moist medium such as clean sand or vermiculite at appropriate temperatures for a specified period of time. The idea is to mimic critical conditions necessary for germination that seeds are exposed to naturally in the environment after dispersal. Generally, if seeds are dispersed in autumn, they may require cold, moist stratification. If dispersed in late spring or early summer, they may require warm, moist conditions, or warm followed by cold stratification, as in Michigan lily (Lilium michiganense). Mix seed with an equal amount of moist, sterile sand, sawdust, peat, or vermiculite and place in a zip-closure bag or small plastic container. Avoid excessive moisture; water should not be pooled anywhere in the bag or container. Use vermiculite if working with species adapted to drier conditions to minimize the risk of rot. If it is desirable to separate the seed from the medium after stratification, one strategy is to place the seed in a small fine-mesh (organza) bag such as those used for wedding favors, then sandwich the bag between layers of moist stratification media.

Effective temperatures for cold stratification are from 32 to 45 °F (0 to 10 °C), with 41 °F (5 °C) considered optimal for many species (Baskin and Baskin 1998). Some species require as little as ten days, others as long as 120 days. A few species, among them American vetch (Vicia americana) and butterfly milkweed (Asclepias tuberosa), will germinate at these temperatures, so check bags weekly to look for emergence of the radicle and plant immediately if this occurs. Some species may germinate best when stratified under natural winter temperature fluctuations (for example, in an unheated building). If sowing seeds in flats for outdoor stratification, cover with screen mesh to protect seeds from being displaced by animals or heavy rains. Cold frames can be used for stratification and extending the growing season in the spring. Effective temperatures for warm stratification are from 68 to 94°F (20 to 35°C), with 68 to 76°F (20 to 24°C) optimal for many species with this requirement (Baskin and Baskin 1998).

Rhizobium Inoculation

Rhizobium are types of nitrogen-fixing bacteria that live symbiotically with the roots of many species, typically forming nodules on rootlets of the plant. They “fix” nitrogen by converting gaseous nitrogen from the air spaces in the soil into plant-available ammonia nitrogen, which directly benefits the host plant. The plant, in turn, provides carbohydrates for the rhizobium. Strains of rhizobium have been isolated and are commercially available for groups of species, notably the genera Amorpha, Lespedeza, and Dalea. Rhizobium comes as a black powder that is mixed with the seed just prior to planting. Greenhouse-grown seedlings of legumes benefit from rhizobium inoculation. It may be unnecessary to inoculate with rhizobium, however, if seedlings will be planted within a few weeks after germination into native soil where rhizobium naturally occurs.

Mycorrhizal Inoculation

Mycorrhiza means “fungus root” and describes a symbiotic relationship between plants and fungi. This is common in many, if not most, plant species. Mycorrhizal fungi are naturally occurring in healthy soil, but may need to be provided for soils that have been fallow, flooded, or eroded over long periods. Sites disturbed by extensive and long-term construction grading or altered by mining will also benefit from inoculation. Commercial inoculum consisting of endomycorrhizal spores, host plant roots, and a sterilized medium is now available, and this can be incorporated into the soil at the time of seeding or transplanting. Inoculation of the site with healthy soil from a different location is another option.

Greenhouse Propagation

Once viable seed is obtained and pre-treated to remove dormancy, it is ready to plant in the greenhouse. Critical factors include a suitable container and potting medium, water, soil temperature, light, and air.

Containers

Containers should provide good drainage and space for root development and yet be small enough to provide efficient use of potting medium and bench space. Nurseries interested in retail seedling/plant production will first germinate seeds in flats, which take up much less greenhouse space. Seedlings are then transplanted into disposable trays with perforated pull-apart planting cells for retail marketing.

Seedlings for transplant into seed production rows can be grown in plug trays or a modular reusable “cone-tainer” and tray system. In recent years, the TPC has had success with 2.5” deep, 73-cell plug flats that are ridged to direct root development downward and have ¾” bottom openings to encourage root pruning and the formation of firmly rooted plugs for transplanting. It is easier to maintain even watering with these plugs than deeper cone-tainers, and a transplantable plug can be produced in a shorter time for most species. In addition, the shallower depth of holes required for transplanting into production rows means that the holes are less apt to collapse and require re-digging. We observe little mortality of these plugs transplanted into prepared fields in spring, generally in May to early June. In very dry years, we provide supplemental water once or twice a week for the first few weeks in the field, but normal rains are usually sufficient for establishment of transplanted plugs.

Images (left to right): seedlings of common boneset (Eupatorium perfoliatum) with first true leaves in a germination tray (with wooden chopstick for scale); one of the seedlings extracted with tweezers to show roots at suitable stage for dibbling into plugs (the chopstick serves to dibble a planting hole in each plug); seedlings in plugs two weeks after dibbling.

In the past, the TPC used Stuewe “cone-tainers” for greenhouse propagation. Cone-tainers are designed for accommodating taproot growth in conifer tree seedlings. They worked well for perennial prairie species – particularly those that put down tap roots like compassplant (Silphium laciniatum), butterfly milkweed (Asclepias tuberosa), and Baptisia spp. Various sizes of cone-tainers are available in yellow UV-stabilized plastic for longer life. The so-called “fir” cells, approximately 6.5 in deep with a 1-in. diameter at the top, work well for most native species. Each tray accommodates 200 cone-tainers, 100 cone-tainers per square foot, so it’s also an efficient use of limited bench space. Planting “dibbles” are available that match the size and shape of the cone-tainers. The cone-tainers and dibble system allow the roots of seedlings to be planted deeply enough so that roots can tap into subsoil moisture. Irrigation isn’t necessary, especially with spring or fall transplanting when rains are more frequent.

There is no single “best” container for prairie plant propagation. Some species that form fleshy basal rosettes in the first year of growth (e.g., Lobelia spp.) seem to benefit from being started in wider, shallower plugs or pots. For species that are sensitive to root disturbance during transplanting, consider producing plugs in biodegradable containers such as coir pellets or Stuewe Zipset Plant Bands, made from milk carton paperboard.

Images (left to right): cardinalflower seedlings (Lobelia cardinalis) nearly one month after dibbling from a germination flat into 6-pack trays; the same seedlings with leafy basal rosettes three weeks later; and plants ready for transplanting to the field six weeks after dibbling into 6-packs.

Potting Medium

A good potting medium should be light enough to allow for good root development, provide adequate drainage, and have enough fertility for seedlings to grow quickly in the greenhouse environment and become large enough for transplanting in a reasonable amount of time. It should also be sterile, meaning free of weed seed and disease organisms. We recommend a soilless mix (less than 20% soil) consisting of 10% sterile soil and 10% composted manure, about 50% milled peat moss or coco coir, and the remainder equal parts perlite and vermiculite.

Caution: All of these materials are extremely dusty. Wear a high-quality particulate mask when handling and mixing materials. Gloves and dust goggles are also recommended. Materials should be moistened before mixing to reduce dust.

Pre-mixed sterile potting mix for plug production can be purchased in bulk from nursery and greenhouse supply companies. Some formulations include potentially beneficial additives such as mycorrhizal inoculum (generally a single species) and biological fungicide (a bacterial species). For propagation of most prairie plants, we supplement “general purpose” mixes by adding additional perlite (2.8 gallons), vermiculite (2.8 gallons), and coated, slow-release fertilizer pellets (19 oz, 15-9-12 formulation) per 3.8 cubic foot compressed bale (7 cu ft expanded). Seedlings are susceptible to damage from over-fertilization, particularly legumes. Controlled-release fertilizer has the added benefit of continuing to provide fertility after transplantation of seedlings into production plots. A drill attachment designed for mixing drywall joint compound helps combine these ingredients while also breaking up chunks of compressed media.

*Image: Small amounts of prepared soilless mixes can be amended and mixed efficiently with a joint compound mixer. Goggles and a particulate mask provide protection from dust particles.*

If you prefer to mix your own potting medium, the recipe below makes approximately 1 cubic yard of soilless mix (enough for 40 trays of 200-cell “fir” cone-tainers). To improve the sustainability of this mix, you could substitute coco coir for the peat moss. Coir is available in compressed, dry bricks that expand greatly when moistened (one 11-lb brick expands to 3 cu ft). If mixing on the floor, sweep and vacuum the area before mixing to remove seeds, debris, and other contaminants:

Peat moss (4 cu. ft/bag) or coir (see above text)	2 bags (8 cu. ft)
Vermiculite (medium texture, 4 cu. ft/bag)	1/2 bag (2 cu. ft)
Perlite (4 cu. ft/bag)	1/2 bag (2 cu. ft)
Sterile soil	two 5-gal buckets
Composted (sterile) manure	manure 40-lb bag
Osmocote® Plus fertilizer 15-9-12 (180 days)	8 lb

Screen peat moss, soil, and composted cow manure through a ½-in. mesh hardware cloth screen to break up or remove large pieces that would tend to clog and create air pockets as cone-tainers are being filled. Add remaining ingredients, mix with shovels on a clean floor, and fill trays.

Strategy for Filling and Seeding Cone-tainers or Plug Trays

We fill several plug trays at a time within a larger, heavy duty garden tray, banging the large tray several times to settle the medium and sprinkling the plugs with water after each addition of soilless mix, until the level of the medium is consistent across the plugs and flush with the top of the plug tray. It will continue to settle somewhat as the trays are watered in, leaving a little space at the top for water to puddle.

A picture of a person leveling off the soil in a set of plug flats. — *Filling plug flats with potting mix.*

When filling trays of cone-tainers, tamp the tray on the floor to firm potting medium and remove large air pockets. Avoid overfilling. Leave about ¾-in. unfilled space at the top of each cone-tainer. This space acts as a reservoir during watering, allowing the water to seep in slowly, helping to saturate the entire soil column. Water cone-tainers frequently for a day or two before planting to fully hydrate the potting medium. Refill any cone-tainers that may have settled excessively.

Attempt to sow several seeds per cone-tainer or plug cell. If seed has been mixed with damp sand or other medium for stratification, it may be impossible to distinguish small seeds from the medium. If this is the case, place the damp seed/medium mixture into a shallow dish and mix thoroughly to evenly distribute the seed within the medium. Use a small, flat implement (point of a knife or wood popsicle stick) to place a small amount of mixture in each cone-tainer.

Getting an appropriate number of seeds per cell is guesswork with tiny seeds, but experience will improve efficiency. Thinning may be necessary if too many seeds germinate in each cell, and this could lead to unintended selection. Blank cells will result if no viable seeds were planted, wasting greenhouse space and making it more difficult to achieve even watering. For larger seeds with high purity and viability (90% or more), one to three seeds per container is adequate. Increase accordingly for seed of poor or unknown quality. Cover with no more than 1/8- to ¼-in. of soil for most species. Caution: Very tiny seeds should not be covered! Additional information and precautions on sowing seed of specific species can be found under the following sections on light, temperature, and water requirements for propagation.

Image: Samples of stratified seed are spread evenly on the surface of prepared germination trays. Small seeds such as these skyblue aster achenes (Symphyotrichum oolentangiense) are very lightly covered with soil.

One strategy to ensure even planting of plug trays is to start seed in germination flats, then prick out seedlings with forceps (tweezers) and transplant them individually into plugs when ready, generally when the first set of true leaves has emerged. There is a risk that individuals with genetic traits that cause them to germinate later could be selected out of the population by this practice. To avoid selection against later germination, retain the germination flats and add later emerging seedlings to plugs when they are ready.

Image: Seedlings from germination trays are dibbled into plugs using forceps, ensuring that all plugs in a flat are filled. This practice is labor-intensive but makes efficient use of seed and greenhouse space.

Light

See Appendix A, Table 2A for recommended planting depth and light requirements

Natural light should be sufficient for seedling establishment in the greenhouse from mid-March through mid-September. Sow seeds in early February and expect germination and emergence to occur over a two- to six-week period. Greenhouse-grown seedlings grow well with only natural light through March and April and into May, when transplanting into production plots begins. Some species require light for germination. These are typically small-seeded species, including Culver’s root (Veronicastrum virginicum), mountainmints (Pycnanthemum spp.), grass-leaved goldenrod (Euthamia graminifolia), bonesets and Joe Pye weeds (Eupatorium and Eutrochium spp.), great blue lobelia (Lobelia siphilitica), and prairie sage (Artemisia ludoviciana). These do best if sprinkled on top of the soil surface and kept continually moist until the seed leaves (cotyledons) are evident.

Temperature

Germination will occur throughout a range of temperatures but will be slower with less than optimal temperatures. The risk of fungal pathogens and rot increases if seed does not germinate and is non-dormant. Warm-season grasses and legumes germinate best in warm soils greater than 70°F (21°C ). Cool-season grasses and many forb species will germinate more readily in cool soil temperatures of 40 to 50°F (5 to 10°C) and may cease germination at temperatures above 77°F (25°C). Soil temperature in the planting trays fluctuates with greenhouse air temperature (72°F, 22.2°C daytime and 60°F, 15.5°C nighttime). Pulses of emergence occur on sunny days with some species, presumably because an optimum soil temperature has been reached from solar heating. Covering plug trays or cone-tainers with translucent plastic increases soil temperature and improves the germination of species that require warm soil temperatures. Use this technique with caution. Lethal temperatures can occur quickly under the plastic with full summer sun. Plastic should be removed as soon as the first seedlings emerge to avoid overheating new seedlings. Cooler soil temperatures can be achieved by setting trays on the floor. If sowing seed in germination flats, precise regulation of soil temperature can be achieved with propagation mats. Propagation mats are placed under the flats and are plugged into an electrical temperature control box. Soil temperature in the flats is regulated by a soil temperature probe from the control box inserted into the potting soil of one of the flats. These are commercially available at reasonable cost from nursery or greenhouse supply companies.

Watering

Proper watering is critical to greenhouse propagation. Watering methods must be adjusted depending on the growth stage of the plants. It is important to keep the soil surface moist until germination has occurred, especially for small seeds that require light and are sown directly on the soil surface. An automated mist system is helpful during this stage of propagation. If using a wand or watering can, use a sprinkler head that produces small gentle droplets and low pressure so that watering doesn’t dislodge seed, force it deeper into the soil, or splatter it out of the containers.

Established plants should be watered thoroughly about once a day, until water drains from the plugs, ensuring that the entire soil column is moistened. Allow the soil to drain and the surface soil to begin to dry slightly between waterings. Shallow, light watering will cause the lower portion of the soil to dry out and root growth to stall. Containers that are overfilled with soil are prone to underwatering since the water can’t pool on the surface and gradually soak in. Overwatering saturates the soil, depriving the root zone of air and creating conditions conducive to decay. Containers and potting medium that allow for proper drainage will help prevent over-watering. Cool, cloudy days reduce watering needs. More frequent watering (two to three times daily) is required on hot, sunny days and for larger plants with fibrous roots that fully occupy the container.

Excessive watering can also damage shoots. Healthy-looking plants will suddenly fall over, appearing to be cut off at the soil level. This is known as “damping off” and is caused by a fungus. Legumes are particularly susceptible to this condition, but it can affect other species as well, especially if they are planted too densely. Sprinkling a layer of fine chick grit (found in the feed section of farm supply stores) over the top of the soil surface after seeding or transplanting will help dry the soil surface and wick water away from the stems. Maintaining good air circulation will evaporate excess water from stems and the soil surface and minimize the risk of damping off. Place additional fans near benches with species known to be susceptible. Thinning may be necessary to improve air circulation. Watering pots/containers from below by setting them in a pan of water until the soil wicks up moisture may also help. Washing and sterilizing containers, benches, and equipment and using sterile potting media also help reduce the risk of damping off.

Image: A topping of chick grit keeps the surface around seedlings dry, helping to prevent outbreaks of damping off fungi.

Transplanting Seedlings

Seedling Development/Timing

The key to successful transplanting of native perennial species is strong root development. Ideally roots should fully occupy the entire soil column so that when the plant is removed from the container, the soil and roots remain intact as a “plug” (i.e., they retain the shape of the container). Grasses and forbs with fibrous roots form beautiful plugs after a few weeks of growth and present little challenge in transplanting. Species with taproots (e.g., Baptisia spp., compassplant, Desmodium spp., butterfly milkweed) develop a thickened fleshy taproot within a few weeks after germination in a greenhouse, but the taproot itself may not be enough to hold the plug intact when transplanting. However, if growth is allowed to continue until the taproot reaches the opening at the bottom of the container, the taproot will air-prune, and fine lateral root growth will be stimulated. These fine lateral roots will help considerably in holding the plug intact when the seedling is removed from the container for transplanting. Slower growing forbs and shrubs require more time for roots to develop adequately for transplant. This process is accelerated by using the 2.5” deep plug trays with ridges and large bottom openings, as opposed to cone-tainers.

Image: Plugs of a taprooted species, longbract wild indigo (Baptisia bracteata, cream wild indigo). Firmly rooted plugs were produced by growing seedlings in air-pruning containers with grooves and large bottom openings.

Seedlings are prone to top-kill when transplanted directly from the greenhouse into the field. Greenhouse-grown seedlings are pampered: they have been protected from drying wind, harsh sun, and herbivores. Robust greenhouse-grown seedlings of many native species can tolerate the stress and will regrow quickly if they have strong root development and adequate soil moisture. A better approach, however, is to acclimate seedlings gradually to outdoor conditions with a process called “hardening off.” A week or two before transplanting, set flats or trays outside for a few hours each day from mid-morning to mid-afternoon in a place sheltered from strong winds and full sun. The idea is to acclimate the plants gradually to outdoor conditions of wind and sun. Strong winds and heavy rains should be avoided. Another option is to move flats/trays into a cold frame (unheated greenhouse) and roll up the sides or open the side vents to allow natural airflow and some direct sun to the plants.

The ideal time for transplanting is in the spring just after the last frost-free date for your region. Rains are more reliable at this time of year, the sun less intense, and plants have the entire growing season to establish and flourish. If transplanting during the summer months, check for adequate soil moisture. Be prepared to water regularly and deeply until plants are established. Transplanting in the fall (early to mid-September) may be another option. Plants transplanted after mid-September may not have enough subsequent root growth to anchor them in the soil, making them vulnerable to frost-heaving.

Bare Soil vs. Weed Barrier

To establish seed production plots, seedlings can be transplanted into weed-free bare soil or a weed barrier. Transplanting into killed sod works well for natural landscaping with a mixed assemblage of prairie species but is not recommended for seed production plots. The dead sod interferes with cultivation and hoeing should weeds become a problem.

Aggressive clonal species like wild bergamot (Monarda fistulosa), prairie coreopsis (Coreopsis palmata), Culver’s root (Veronicastrum virginicum), and prairie cordgrass (Spartina pectinata) can be successfully established as seed production plots in bare soil. Good weed suppression, as always, is critical. Pre-emergent herbicide (Pendulum®, Prowl®) can be used on the site after transplanting to inhibit weed seed germination. Be sure to water in seedlings after transplanting to settle soil around the root zone before applying pre-emergent. Otherwise rain may concentrate run-off of herbicide into freshly dibbled holes containing the new transplants, damaging roots or killing the seedlings. Do a small test plot on species not listed as tolerant on the product label. Read, understand, and follow all precautions and directions on the product label. Planting 8 to 12 in. apart in well-spaced rows (32 to 36 in.) will permit cultivation later that season for weed control. Do not cultivate as long as pre-emergent herbicide remains effective in suppressing weed seed germination. These aggressive clonal species will spread and create solid rows in two to three years. Seed production begins to decline after the third year and may drop off sharply after the fourth year as plants become root-bound within the rows. Aerating the soil with a turf-type aerating implement in the fall or early spring may extend plot productivity.

Weed barrier comes in a variety of materials: plastic film, woven fabric, paper, and fiber mats. Plastic film is inexpensive but less durable than most other types of barriers. Plastic creates abnormally moist conditions that can increase disease problems in some species but may be suitable for short-term (one to two-year) applications. Biodegradable plastics are available which begin breaking down after the first growing season. Plastic film can be applied mechanically using an implement that creates a flat or raised bed while simultaneously rolling out and securing the plastic. Irrigation tubing can be laid at the same time for species needing additional moisture. Plugs can be transplanted into holes dibbled through the plastic film. When using 4-ft wide plastic, we typically plant four plugs per row in rows about a foot apart. We adjust this plant spacing to accommodate larger species such as compassplant.

Image: Metal dibbles punch planting holes directly through plastic weed barrier. Most species are planted in rows of four spaced one foot apart.

As the name implies, a weed barrier suppresses weed growth by blocking light, physically obstructing shoot growth, and by solar heat sterilization of weed seeds in the soil. Weed barriers also conserve soil moisture, and plants grow more vigorously with a weed barrier than in bare soil. For these reasons, weed barriers aid establishment in the first year and increase seed production in the second year for most species. By the third year, aggressive clonal species will send rhizomes in all directions that the barrier would smother. For these species, plant in bare soil as described above, or use an inexpensive barrier that biodegrades or can be pulled up after the second season.

In the past, we used a durable, water- and gas-permeable weed barrier (DeWitt Pro woven landscape fabric) for long-term applications (up to ten years). This can be purchased in rolls 6’ x 250’ or 12’ x 300’ rolls. This type of weed barrier provides long-term benefits of weed suppression and moisture retention for less-competitive species that establish slowly. It’s also a good choice for species with taproots that remain where they are planted (non-clonal). However, removal and disposal of fabric weed barriers after many years in the field is very difficult, as perennial grasses and other weeds grow through the degrading barrier, uniting it with the soil.

Transplant seedlings of non-clonal species into weed barrier fabric 8 to 12 in. apart in blocks or rows to optimize the use of this costly product. Slits are cut in the fabric weed barrier using a tool made of modified sickle bar blades mounted on a wooden 2x4 with an attached handle. This device cuts four “x-shaped” 2-in. slits, 8 in. apart at one time. Cutting larger slits would allow more weeds to germinate and come up through the openings. Precise holes can also be burned into weed barrier fabric using a propane-fired hole cutter available from greenhouse and nursery supply companies.

Regardless of the type of barrier used, some hand weeding will be necessary, especially during the first growing season. Hand-pull weeds that emerge through the barrier openings while they are small, taking great care not to disturb new transplants. Pressing gently downward on the top of the newly transplanted plug while pulling adjacent weeds will protect it from being uprooted. A garden or dandelion knife can be used carefully to slice through the taproots of larger weeds. Slice into a taproot just below the soil surface, again taking care not to damage the roots of transplants. Hand weeding periodically throughout the first growing season will be required to give transplants the best chance of establishing the first year and producing a good seed crop in subsequent years.

How to Transplant Seedlings

Soil should be firm in all cases. If dibbling into bare soil, the soil should be rolled or packed to prevent dibbled holes from collapsing. Likewise, very dry soils resist dibbling. It may be necessary to sprinkle the area the day before or wait until a day or two after a soaking rain. Just before transplanting, liberally water seedlings to fully saturate the root plugs. This will make it much easier to remove the plugs from the trays or cone-tainers and provide extra moisture to the root zone after transplanting. To remove plants from a plug tray, use a stick or gloved finger to push up through the opening at the base of each plug. Hold the plant at its base and pull gently. If the plant does not release easily, try “massaging” to loosen the root plug from the sides of the cell. If the plants are in cone-tainers, hold a cone-tainer upside-down firmly in one hand, and rap the rim of the cone-tainer sharply on the palm of the other hand, using a flick of the wrist. The plug should slide out easily; repeat if necessary. (If plugs do not hold their shape upon removal, either the roots are not adequately developed or too much force is being used. Transplant success drops significantly if this happens!) Slide the intact plug into a dibbled hole – it should just fit, with the top of the root plug just at or slightly below the soil level. Pinch soil firmly around the top of the plug to seal in moisture, taking care not to bury the base of the shoot. The lighter soil-less mix can wick moisture away from the roots if left exposed. Be sure dibbled holes are deep enough to comfortably receive the full depth of the plug. Plugs forced into a too-small or shallow hole will often pop out of the ground after a good rain, exposing the root collar.

Image (left to right): Well-rooted plugs pop out of trays intact; plugs are "pinched in" to achieve good contact between roots and soil and topped with a thin layer of field soil to prevent moisture from wicking out of the potting mix.

References

Baskin, C. C., & Baskin, J. M. (2003). When breaking seed dormancy is a problem: Try a move-along experiment. Native Plants Journal, 4(1), 17–21. https://doi.org/10.3368/npj.4.1.17

Baskin, Carol C., & Baskin, J. M. (2014). Seeds: Ecology, biogeography, and evolution of dormancy and germination. Elsevier / Academic Press.

Deno, N. C. (1993). Seed germination: Theory and practice. available for free download from the USDA National Agricultural Library at https://search.nal.usda.gov/permalink/01NAL_INST/27vehl/alma9916347016207426

Khadduri, N. Y., & Harrington, J. T. (2002). Shaken, not stirred - a percussion scarification technique. Native Plants Journal, 3(1), 65–66. https://doi.org/10.3368/npj.3.1.65

Native Plant Network, USDA Forest Service. (n.d.). Propagation protocols database. Reforestation, Nurseries, & Genetic Resources (RNGR). https://npn.rngr.net/propagation/protocols

Society for Ecological Restoration and Royal Botanic Gardens Kew. (n.d.). Seed information database. https://ser-sid.org/about

Harvesting Native Seed

Harvesting Native Seed sagem Wed, 01/17/2024 - 12:30

Proper timing of seed harvest and use of effective harvest techniques increases the likelihood of producing a profitable yield of viable seed that will be effective for restoration of native vegetation. Harvest is clearly a critical step in seed production. It is also one of the steps that is most susceptible to unintended selection leading to adaptation of wild species to production systems. Wild populations have variation in inherited traits related to seed harvest such as the timing of maturation and the degree to which seed shatters or is retained on the plant at maturity. Variation in these traits provides wild populations with the ability to adapt to changing conditions and to disperse and establish without human intervention. Loss of these traits within production populations is possible when all seed in a field is harvested at peak maturity (eliminating the genetics of earlier and later maturing individuals from the harvest) or when most of the harvested seed comes from individuals that retain seed longer (i.e. are less prone to shattering). The risk of selection is compounded as harvested seed is used in expanding fields and/or planting new fields over several generations. Keeping the risk of selection in mind and utilizing strategies to avoid it is key to ensuring that commercially produced seeds retain genetic diversity and the ability to grow, survive, and reproduce as wild plants in restoration sites.

Gauging Seed Maturity

Factors Affecting Seed Maturity and Dispersal

Gauging seed maturity is key to harvesting quality seed. Immature seed stores poorly, losing viability more quickly than mature seed. Determining the ideal time to harvest can be difficult. Within a population of native plants, there is genetic variation among individuals in the timing of seed ripening, and repeated harvests may be called for as seeds mature on different plants. In addition, seeds in different parts of an individual seedhead or inflorescence are often at different stages of ripeness. Seed maturity usually progresses from top to bottom of the seedhead in grasses and many prairie forbs. In forb species with flattened flower clusters, seed often matures first at the center of the inflorescence. Mature seeds may be quickly dispersed by gravity, wind, water, or animals, so it’s important not to delay harvesting. Some species forcefully eject seed at maturity (e.g., spiderworts, phlox, wild petunia, and violets) and require special harvesting strategies.

A conceptual graph illustrating the maturation of seed and seed shattering with an estimate of the optimal harvest time — Conceptual diagram illustrating the optimal harvest time for capturing maximum mature seed yield, assuming the field is harvested mechanically on a single day. As seed ripens, seed shattering follows, and the rate of both processes increases over time. Timing the harvest carefully maximizes seed yield and quality.

Environmental factors also influence seed maturation. Cold, moist conditions tend to delay seed maturity while hot, dry conditions may hasten it. Latitude also affects ripening since many plants flower and set seed in response to photoperiod. Flowering and seed set may be delayed if plants are grown northward from their origin, or hastened if moved southward. If moved a great enough distance north or south of their origin (greater than 300 miles) they may fail to set mature seed.

Three fields of butterfly milkweed sourced from different zones photographed on the same day. — Three fields of butterfly milkweed (Asclepias tuberosus) showing differences in flowering phenology, photographed on July 3, 2023 at the Tallgrass Prairie Center, UNI. The northern Iowa zone (left) is nearly finished flowering, central Iowa zone (center) is just past peak flowering, and southern Iowa zone (right) is approaching peak flowering.

Gauging Seed Ripening in Grasses

Seed ripening and timing of harvest varies by species, source of parent material, and environmental conditions. For example, cool-season grasses begin growth early in the growing season and consequently ripen earlier as compared to warm-season grasses. In grasses, there are roughly four stages of seed maturity: milk, soft dough, medium dough, and hard dough. Firm thumbnail pressure on the caryopsis, or grain, will help determine maturity. Grasses should be harvested at the hard dough stage, when firm thumbnail pressure slightly dents the caryopsis. Many grasses ripen about a month after flowering, and some do not hold seed long after maturity. Test ripeness by firmly striking the seed head against your palm; if some shattering occurs, the seed is ready to harvest. If it shatters with only gentle striking, harvest immediately. In some species, such as sideoats grama, visual assessment of the stand provides an indicator of maturity: the stand is ready to harvest when about 10% of the plants have started to shed seed.

Gauging Seed Maturity in Forbs

Generally, the seedhead itself or the stalk immediately below the seed head will begin to appear dry or discolored as the seed ripens. The leaves may also change color as the plant approaches dormancy. If the seed is easy to strip off or shatters out when the seed head is gently struck against the palm, seed ripening has begun. Some species have seed capsules that dry and split open when ripe, allowing seed to be released by wind, passing animals, or gravity. The spiderworts (Tradescantia), members of the day-flower family (Commelinaceae), are a special case: they drop seed from individual seed capsules as they ripen even while the bracts remain green and other flowers in the same cluster are in bud or blooming. Species with dispersal apparatus such as awns or hairs will appear dry and fluffy at maturity. It can be tricky to time the harvest of fluffy-seeded species that release their seed soon after maturity (many species in the aster family). Picking a few heads and examining the developing “seeds” (technically a type of fruit called cypselae or achenes) can help gauge approaching maturity. Seed that is nearly ripe will develop the mature color and separate easily from the receptacle.

A plant with gray seedheads and brown, dry stalks indicating the seeds are ready to harvest — *A hairy mountainmint plant (Pycnanthemum pilosum) showing signs that it is ready to harvest: leaves are changing color and dropping and stalks beneath the seedheads are dry and brown.*

Open seed capsules of three forb species — Three species of forbs showing capsules that have opened and begun releasing seed; they should be harvested immediately to avoid losing the crop. Left to right: bottle gentian (Gentiana andrewsii), swamp lousewort (Pedicularis lanceolata), and cardinalflower (Lobelia cardinalis).

Collage of images showing different stages of seed ripeness in butterfly milkweed — Fruit and seed ripeness varies within a seed production plot. Left: a field of butterfly milkweed with "pods" (follicles) at varying stages of ripeness. Center: Once pods split naturally, the seed rapidly blows away. Right: A pod that bursts in response to gentle pressure on the seam reveals chocolate-brown, mature seeds; this is the ideal time to harvest.

Aster seed heads showing characteristics of ripeness: dry fluffy pappus and changing color of seeds. — Maturing seedheads on a sky blue aster plant (Symphyotrichum oolentangiense). The greenish receptacle near the center shows three indicators of approaching seed maturity in many members of the aster family: fluffy pappus, mature color of achenes, and easy separation from the receptacle.

Hand Harvesting Techniques

Hand harvesting is time- and labor-intensive and not practical for most large projects, but it is an important way to collect the seed of native species that are commercially unavailable and/or impractical to harvest mechanically. Such species may be low or high growing species, early or late ripening species, species with explosive seed capsules, or uncommon or patchy species in native prairie. There are also situations in a seed production system when hand harvesting may be needed, for instance, when collecting seed from early maturing individuals to ensure conservation of genetic diversity. Efficiency can be improved by keeping both hands free to harvest by simply fastening collection bags and containers around the waist with a bungee cord. Hand harvesting aprons are also available from some native seed businesses and garden suppliers. A plastic comb can be used to strip seed from grass stems. Scissors, clippers, or a hand sickle are useful for cutting tough stalks beneath seed heads.

Three situations in which hand harvesting is warranted — Three situations in which hand harvesting was warranted: Left - stripping seed from sedge plants in a plot that was partially invaded by another sedge species (note hands free technique); Center - using a sickle to cut early maturing individuals in a prairie Junegrass plot (the plot was later combined); Right - hand stripping sticky Illinois ticktrefoil pods which tend to ball up in the combine (gloves and picking apron are shown; raincoat and rainpants, not shown, help to protect clothing).

Hand harvesting or combinations of mechanical and hand harvest approaches may be needed for species with explosive seed capsules or capsules that shed seed immediately upon ripening. Strategies for collecting explosive seeds include: checking and harvesting daily (Viola), bagging seedheads loosely with a tightly woven mesh or cloth bag (Phlox), vacuuming seed from landscape fabric (Phlox), or clipping heads when some capsules have opened and drying cut material on tarps or cloth bags indoors (Ruellia). For the spiderworts (Tradescantia), check seedheads regularly near the typical harvest date. They begin shedding seed from the bottom of a flower cluster even when most of the plant is still green and there are still a few buds opening at the top of the cluster. When some seed is being shed from most plants, plots can be swathed and the cut material collected and laid on a tarp in a building. Capsules will continue to mature and release seed as the cut material dries.

Three pictures showing how to harvest mature seed of longbract spiderwort — Longbract spiderwort (Tradescantia bracteata) seed capsules mature and release seed even while other buds in a cluster are still flowering. Left: examining the underside of a seed cluster for open capsules. Center: hand cutting spiderwort stalks (the plot could also be swathed mechanically). Right: cut stalks drying in a building. The tarp catches the seed as the capsules mature and break open.

Hand harvesting is also useful for collecting seed from wild populations for use as foundation material for seed increase and seed production plots. A seed collection is only a sample of the seed available in a population. How and when the seed is collected and propagated influences the genetic potential of the resulting population. Annual variation in rainfall and temperature can affect total seed production, quality, maturity, and dormancy. To optimize the capture of genetic diversity present in a population, approximately equal amounts of seed should be collected from many widely spaced individuals (minimum of 30, preferably 50 or more) throughout a site and over multiple years. In larger, evenly distributed populations, walk a linear transect through the population, sampling seed at intervals (e.g., every ten steps), then sample along additional transects parallel to the first. When collections from multiple sites are used to create a pooled population in a production plot, attempt to equalize the contribution of seed from each site.

Two people harvesting seed from a remnant prairie in fall — *Individual team members can collect along parallel transects to capture as much diversity as possible from a large plant population in a remnant prairie.*

Populations grown and re-grown in a production field can become adapted to site conditions and nursery management practices. Even though most prairie plants are perennial, seed production in plots tends to decline after 3-5 years for most species, and plots need to be replanted. Therefore, it is important to save seed from the original collections and/or the first production generation for replanting production fields. Hand harvesting of early maturing individuals in a production plot is a way to retain important genetic diversity within the production population. The rest of the plants in the production plot can then be mechanically harvested at peak maturity. Later maturing genotypes can be retained by leaving a portion of the plot uncut and harvesting it later (either mechanically or by hand).

Mechanical Harvesting Techniques

Combine harvest

As stated above, not all seeds ripen at the same time; with any species, a determination has to be made as to when most seeds present are at or nearing maturity. Grasses are generally harvested at the hard dough stage with some exceptions. Test ripeness by firmly striking the seed head against your palm; if shattering occurs with only gentle striking, the stand should be combined immediately. Since most grasses ripen from the top down, some shattering of the tops of the seed heads may have already occurred. If a species’ seed shatters very easily, harvest in the early morning when humidity is high and wind speeds are less, as a strong wind can reduce the harvest significantly in a single afternoon. In grasses that shatter easily at maturity, windrowing or swathing during the medium- to hard-dough stage can be effective, since seed will after-ripen for several days after cutting. It is important to be sure no rains are in the forecast for the next few days after cutting. Swaths can then be picked up with a combine after the material has dried at the site for a few days.

Combines may require significant modifications to make them suitable for harvest of native grasses and forbs. Reducing or shutting off airflow in a combine is a must for native species with lightweight seeds. Species with fluffy pappus (e.g., asters and goldenrods) should be harvested when seed is mature but just prior to the dry-fluff stage. If the seed is dry-fluffy, the combine will become a super seed dispersal machine. Plugging of shaker sieves and augers is a constant issue, particularly with fluffy seed or seed with long awns. The long twisty awns of Canada wildrye can be a combine’s bane. De-awner bars can be installed into the cylinder surrounding the concaves to increase the threshing action of the concaves. Seed of tick trefoils (Desmodium species) is dispersed inside pods with sticky hairs. Some of the pods ball up in the combine, fail to pass through the sieves, and are ejected from the straw-walker. To prevent significant loss of seed, attach a bin or tarp to catch the seed balls. Alternatively, cut or swath the plot and collect the seed stalks, then pass them through a stationary combine, where the ejected material can be more easily captured.

Combining a native forb plot — *Combining a plot of stiff goldenrod (Solidago rigida). For this and other fluffy seeded species, the airflow must be reduced or shut off.*

Seed strippers

Commercial seed strippers are available as handheld, implement pull-type, or tractor-mounted equipment. They all use a rotating brush or bristles to strip the seed from stems and stalks of plants. While perhaps not as efficient as combines, strippers can be used for the successive harvest of species that ripen gradually or at different times. Handheld strippers and pull-types light enough to be pulled with an ATV allow harvest of otherwise inaccessible native sites.

Leaf blower vacuum units

A leaf blower-vac unit on the vacuum setting can be used to harvest fluffy seed such as species in the aster family that have a pappus (“parachute”). Some suggest that the impeller blades of the machine need to be removed to avoid damaging the seed. However, we carried out a trial using an unmodified blower/shredder vacuum to harvest seed of prairie groundsel (Packera plattensis) and observed no visible damage to the seeds (cypselae) under a dissecting microscope. Because vacuuming removes seeds at the dry-fluff stage, immature seed can remain on the plants to ripen for later harvest. This should increase the yield of viable seed from each plant and reduce the risk of unintentionally selecting for a narrower seed ripening window. An additional benefit of this harvest method is that it removes much of the pappus from the seed at the harvest step, reducing the time needed in the seed cleaning lab. It would be interesting to try scaling up this technique using a modified pull-behind leaf vacuum.

Ethics of Harvesting from Remnant Sites

Repeated, annual harvest of seed from remnant prairies for the commercial market is not encouraged. First, seed production and quality from wild populations are not as high as from production plots, and second, it provides an incentive to manage remnants for maximal seed production. Manipulation of a remnant prairie to maximize seed production – such as whole-site, repeated annual burns, herbicide treatments, or fertilizing – is inappropriate and unethical. A remnant prairie is a diverse, biotic community (both above and below ground) of microbes, fungi, plants, and vertebrate and invertebrate animals interacting in complex relationships. Management applied indiscriminately and repeatedly is detrimental to some of these associations. Burning should be limited to a portion of a remnant any given year, and each portion should be burned on rotation and at different times of year, at varying intervals of time. Bulk harvest from remnants may be appropriate when the seed is intended for planting on adjacent or nearby land to buffer and expand the native prairie. Finally, mechanical harvest in remnant sites should include a careful inspection and cleaning of equipment (including vehicles) to avoid introducing exotic/invasive species that may lead to the degradation of the remnant.

Be mindful of other ethical considerations when collecting seed from prairie remnants. Federal and state endangered and threatened species cannot be collected without proper permits. Removal of any plant or plant part from preserves, natural areas, and parks may be restricted. Check with the proper agency before harvesting seed in these areas. Harvesting from roadsides may also be restricted in some states and counties. Contact the county engineer’s office or state department of transportation before harvesting from county and state roadsides. Obtain permission from the landowner before collecting seed on private property. Some seed producers lease native prairies to compensate landowners for excluding grazing over the summer so that seed can be harvested from the site.

Entrance sign at Clay Prairie Preserve — Obtain proper permits and permissions before making collections on private or public lands. Determining which entity owns or manages a site can be challenging. In the case of Clay Prairie, the land is owned by UNI, but the county helps manage the site, and the prairie is part of the State Preserves system.

Seed Processing

Seed Processing sagem Wed, 01/17/2024 - 12:31

This section is under construction and new material will be added as soon as it is complete.

Post-harvest processes include drying, pre-cleaning, cleaning, and proper seed storage. The extent of processing required after drying depends on the species, storage conditions, and intended seeding method. Knowledge, skill, and access to specialized equipment are necessary for some of the cleaning steps described, and these factors will determine the quality of the finished product. Most commercial producers of native seed clean seed to a very high quality of purity and germination.

As in other steps in native seed production, care must be taken to avoid selection and loss of genetic diversity in production populations. In seed processing, there is a risk of selecting against genes that influence seed traits. For example, if only average-sized seeds are kept and used for expanding or re-growing fields, genes for seeds that are smaller or larger than average can decline in the population within a few generations.

Drying

Drying is a necessary step for proper seed cleaning and storage for nearly all prairie species. Seed should be dried immediately after harvesting to prevent fungal growth and decomposition. Plastic bags or airtight containers can be used for storage ONLY after the seed is properly dried and cleaned. Larger quantities of material require specially constructed bins with screened bottoms and a source of airflow up through the material. Smaller quantities can be collected in large 100% cotton muslin bags made to fit inside a 30-gal plastic trash bin. Large cotton laundry bags with drawstrings work well. Fill bags loosely with seedheads, tie closed, label, and place in drying bins. If the material associated with the seed is very green, as is the case with spiderwort (Tradescantia spp.), or is damp from a recent rain shower, spread the material out on tarps and position several box fans overhead, turning the seed frequently with pitchforks, rakes, or shovels. Drying may take several days to a few weeks, depending on quantity of material and drying conditions (ambient humidity).

Green stalks of Tradescantia laid to dry on a tarp — Image 2: Cut stalks of *Tradescantia bracteata* laid to dry on a tarp.

Pre-cleaning

Harvested material requires some degree of pre-cleaning to reduce bulk. The extent of pre-cleaning required depends on the method of seed harvest, intended storage period, and method of planting. Threshing, debearding/deawning, and brushing are considered pre-cleaning steps, because they are designed to prepare the seed for later cleaning processes by removing unnecessary appendages and improving seed flow.

Threshing

Threshing removes the seed from seedheads, one of the primary functions of a combine harvester. Hand-collected material can be threshed with a variety of machines, including hammer mills, huller/scarifiers, and brush machines. Some growers have adapted older combines (with cutter bars and reel removed for safety) for use as a stationary thresher of hand-collected material. A debearder (see below) is effective for removing the awns of grass seeds and threshing the dried seedheads of compassplant and dried milkweed pods.

A student worker loading milkweed pods into the hopper of a debearding machine — Image 3: Adding dried milkweed pods to the hopper of the Westrup debearder.

Crushed milkweed pods and fluff emerging from a debearding machine — Image 4: Seed falls into a tub positioned below the debearder, mixed with chopped pods and fluff.

Non-mechanized threshing can be accomplished by stomping on seedheads. Using large plastic tubs, place about a 2-in. layer of bulk material in the bottom and stomp on it with waffle-sole boots. Toe kicks to the corners of the tub help break up any stubborn seedheads. Stomped material is then screened through a coarse ½-in. or ¼-in. screen into a second tub. Continue in batches, returning any intact seedheads remaining to the stomping tub. This method is very effective on species of Baptisia, Silphium, Helianthus, Veronicastrum, and Rudbeckia. Echinacea tends to be stubborn and requires machine threshing, unless it is collected late in the season after the seedheads naturally begin to shed their seed.

Deawning/Debearding

Many species have seed appendages such as the awns or “beards” on grasses and pappus “parachutes” on seeds of asters, goldenrods, and other composites. Removing or minimizing these appendages reduces bulk and improves seed flow for further cleaning. This step requires specialized and costly equipment. Debearding machines consist of a rotating shaft with projecting metal bars housed in a chamber. As the chamber fills with seed, the bars work the seed against itself, breaking or rubbing off the awns. It is important to fill the chamber with the proper amount of seed: too little and it’s ineffective, too much and the seed can heat up and be damaged. A continuous flow debearder works well for larger quantities of seed, but batch debearders are available for smaller quantities. A small gallon-size huller/scarifier is useful for deawning small individual accessions of seed. This type of machine is very aggressive, and only a few seconds of treatment are typically needed.

A debearder machine for removing appendages from seed — Image 5: A debearder being used to improve the flowability of combine harvested big bluestem (*Andropogon gerardii*) before cleaning the seed.

A brush machine is useful for removing the pappus from asters, goldenrods, and blazing stars, but this versatile machine has many uses. Its basic action is to use rotating brushes to rub seeds or seedheads over a cylindrical drum screen, or mantle. Mantles come in various screen sizes, and a variety of brush textures is available. The machine can also be used as a deawner or scarifier. It is effective in removing the “cotton” from thimbleweed (Anemone virginiana and A. cylindrica), threshing seed from hand-collected mint seedheads, removing pods from purple prairie clover and leadplant, and deawning smaller quantities of grasses. Heavy canvas beater bars can be installed in place of brushes for a hammer-mill-like effect for removing seed from tough legume pods (e.g., Amorpha spp., Dalea spp., and Astragalus spp.) and dried rosehips (Rosa spp.). Beater bars are constructed from rubber/canvas multi-ply conveyer belting which can be purchased from industrial supply companies and cut to fit. When setting up the machine, position the beaters so that the canvas is barely touching the edge of the screen.

a brush machine with the cover removed to show the screen mantle and brushes — Image 6: A laboratory scale debearder with the front cover removed to show the screen mantle and rotating brushes.

A student worker loading rosehips into the brush machine — Image 7: A student worker feeding dried rosehips into the brush machine which has been set up with canvas beater bars.

Crushed rosehips from the brush machine — Image 8: The brush machine with beaters installed crushes the rosehips and releases the seeds.

Cleaning

Storage

Seed Source and Quality

Seed Source and Quality sagem Wed, 01/17/2024 - 12:32

This section is under construction and new material will be added as soon as it is complete.

Importance of Seed Source
Local Ecotype vs. Regional Sources
Source-Identified Seed
Cultivated Varieties of Native Species
Seed Quality
Calculating Pure Live Seed

Species Production Guides

Species Production Guides sagem Wed, 01/17/2024 - 12:18

This section consists of specific information on seed production for selected species in the following functional groups: forbs (wildflowers), legumes, warm season grasses, cool season grasses, sedges, and woody species (prairie shrubs). Each species production guide includes a description of the species (with photos), its habitat, range map, and conservation status.

Recommendations for stand establishment, management, seed harvest, and seed cleaning are provided. These are based primarily on direct experience at the Tallgrass Prairie Center, together with published information from the NRCS Plant Materials Program, scientific publications, personal communication with other native seed growers, and publicly available databases such as the Seed Information Database and the Propagation Protocols Database of the Native Plant Network. A graph of potential seed yields is provided in each guide, based on harvest records from seed production plots at the Tallgrass Prairie Center.

Each species guide is provided as both a webpage and a printable file (pdf). If you prefer to work with a print version, we encourage you to print pdfs of the species you need and organize them in a binder.

This section is a work in progress. We will continue to add new species guides as they are completed. If you have comments, questions or would like to share propagation information for potential inclusion in these guides, please email laura.walter@uni.edu.

Forbs

Forbs sagem Wed, 01/17/2024 - 12:35

The Species Production Guides for forbs (wildflowers) provide specific information about growing each of these species for seed production.

Scroll the list (alphabetized by scientific name) or press ctrl-f (or command-f) to search for any name in this page.

A printable file (pdf) is also provided on each species page for those needing a print version.

This section is a work in progress. We will continue to add new species guides as they are completed.

Anemone canadensis / Canadian anemone
Anemone cylindrica / candle anemone
Anemone virginiana / tall thimbleweed
Artemisia ludoviciana / white sagebrush
Asclepias incarnata / swamp milkweed
Asclepias syriaca / common milkweed
Asclepias tuberosa / butterfly milkweed
Asclepias verticillata / whorled milkweed
Chelone glabra / white turtlehead
Coreopsis palmata / stiff tickseed
Coreopsis tripteris / tall tickseed
Doellingeria umbellata / parasol whitetop
Potentilla arguta / tall cinquefoil
Echinacea pallida / pale purple coneflower
Eryngium yuccifolium / button eryngo
Eupatorium perfoliatum / common boneset
Euthamia graminifolia / flat-top goldentop
Eutrochium maculatum / spotted joe pye weed
Fragaria virginiana / Virginia strawberry
Gentiana alba / plain gentian
Gentiana andrewsii / closed bottle gentian
Heliopsis helianthoides / smooth oxeye
Helianthus pauciflorus / stiff sunflower
Hypericum ascyron / great St. Johnswort
Liatris aspera / tall blazing star
Liatris ligulistylis / Rocky Mountain blazing star
Liatris pycnostachya / prairie blazing star
Lobelia cardinalis / cardinalflower
Lobelia siphilitica / great blue lobelia
Lythrum alatum / winged lythrum
Monarda fistulosa / wild bergamot

Monarda punctata / spotted beebalm
Oligoneuron riddellii / Riddell's goldenrod
Oligoneuron rigidum / stiff goldenrod
Parthenium integrifolium / wild quinine
Pedicularis lanceolata / swamp lousewort
Prunella vulgaris ssp. lanceolata / lance selfheal
Pycnanthemum verticillatum var. pilosum / whorled mountainmint
Pycnanthemum tenuifolium / narrowleaf mountainmint
Pycnanthemum virginianum / Virginia mountainmint
Ratibida pinnata / pinnate prairie coneflower
Rudbeckia subtomentosa / sweet coneflower
Ruellia humilis / hairy wild petunia
Silphium integrifolium / wholeleaf rosinweed
Silphium laciniatum / compassplant
Solidago missouriensis / Missouri goldenrod
Solidago speciosa / showy goldenrod
Symphyotrichum laeve / smooth blue aster
Symphyotrichum novae-angliae / New England aster
Symphyotrichum oolentangiense / skyblue aster
Symphyotrichum praealtum / willowleaf aster
Tradescantia bracteata / longbract spiderwort
Tradescantia ohiensis / bluejacket
Vernonia baldwinii / Baldwin's ironweed
Vernonia fasciculata / prairie ironweed
Verbena hastata / swamp verbena
Veronicastrum virginicum / Culver's root
Zizia aptera / meadow zizia
Zizia aurea / golden zizia

Virginia strawberry

Virginia strawberry dickeye Mon, 02/03/2025 - 13:27

Virginia strawberry header image

Fragaria virginiana Duchesne

Alternate Common Names: wild strawberry, common strawberry

Scientific Synonyms: Fragaria australis, Fragaria canadensis, Fragaria grayana, Fragaria terrae-novae

Family: ‌rose family (Rosaceae)

Functional Group: forbs (wildflowers)

‌Printable PDF Fragaria virginiana

Description

Life cycle and growth form: perennial, spreading by runners to form extensive clones.
Height: 4-8 in
Leaves and stem: leaves basal, palmately compound; oval shaped leaflets (3) are up to 2.5 in long and 1.5 in wide with coarsely toothed margins and often finely haired, especially on the undersides; stems are sprawling stolons, often reddish in color, that root at the tips where new plants can emerge.
Flower, fruit and seedhead: Flower: regular, five-petaled, 1/2 in diameter, petals white, stamens and pistils in center of flower yellow, 4-6 flowers in a loose cluster that is usually shorter than the leaves.
Fruit/seedhead: 1/2 in globular to ovoid “berry” with numerous achenes (“seeds”) in pits on the berry’s surface, bright red at maturity.
Pollination: Insects such as bees, flies, and butterflies.
Seed: Appendix D
‌Seed characteristics
Seed weight:
Seeds per ounce: 83,000 (Prairie Moon)
1000 seed weight: 0.46g (Seed Information Database)
Description: “Seed” is a reddish-brown achene, 1.2-1.8 mm in diameter, roughly egg-shaped.
Typical seed test
PLS: 83.35%
Purity: 99.81%
Germination: 21%
Dormancy: 61%
TZ: 85%
(averages obtained from 1 test of a purchased seed lot, and 1 test of seed produced at TPC)
Habitat and range: ‌Habitat: Dry to moist soil; partial to full sun; prairies, meadows, woodland openings and edges, roadsides, along railroads, savannas, limestone glades; Wetland Indicator Status is Facultative Upland (FACU) for the Midwest.
Conservation status: Global- G5, secure; Louisiana- S1, critically imperiled; Nevada- S2, imperiled; Illinois- S2/S3, imperiled to vulnerable (NatureServe)

General Comments

Virginia strawberry can be found in the “understory” layer of most remnant prairies in Iowa. These small plants pack a large ecological punch. They flower early in the season, providing nectar and pollen resources for many species of small bees, flies, and skipper butterflies. Various herbivorous insects and mammals feed on the leaves, and the fruits are eaten by birds, mammals, and even reptiles (turtles), which disperse the seeds. When wild strawberries were abundant across the Iowa landscape prior to agricultural conversion, the fruits were an important early summer food for both Native people and Euro-American settlers. Virginia strawberry is one of the species that produced the cultivated strawberry through hybridization. Propagation and processing of this species is not difficult compared to other prairie forbs, and the most significant barrier to seed availability is probably the cost of labor required to pick the berries several times during the 2-3 week fruiting season.

Recommendations for Seed Production

Establishment for seed production

‌Direct seeding

We do not have experience with direct seeding this species for seed production.

‌‌Greenhouse

Seed pre-treatment: 8 weeks cold/moist stratification

Sowing: Sow seed in greenhouse 8-10 weeks before average frost free date; spread stratified seed on germination flats and lightly cover (1/8 in or less) with germination mix; move seedlings to plugs when they have a pair of well-developed true leaves.

Transplanting: When plugs are well rooted and danger of frost is past, harden off seedlings and transplant into weed-free beds prepared with biodegradable paper mulch weighted with clean straw. Mulch must be biodegradable so that the new plantlets that form at the ends of stolons can grow roots into the soil.

‌Appendix A

Stand management

Weeds: Biodegradable mulch in the first growing season suppresses most weeds; mowing at the highest setting of a typical riding mower after the fruiting season keeps weeds from becoming overly competitive in the second year.

Pests: Mammalian herbivores may browse foliage, and birds and small mammals consume fruits.

Diseases: None noted, though diseases that affect commercially grown strawberries such as botrytis molds may become a problem.

Plot renewal: Commercially grown strawberries are known to produce more fruit at the edges of plots. Since wild strawberries are closely related to the cultivated varieties, they may behave similarly, and techniques of cultivation, fertilization, and irrigation that are used by commercial strawberry farms may be beneficial. We will update this information as we gain more experience.

Seed production

First Harvest: Plants flower and set fruit in the second year after planting.

Yield/Acre: 17-18 lbs of seed per acre (extrapolated from first year’s production of one production plot); it takes roughly 30 pounds of berries to obtain a pound of seed.

Stand Life: Probably long-lived, but production may decline as plots grow densely in subsequent years; plants at the edges of clones may set more fruit than those within dense patches; techniques used for “renewing” cultivated strawberry production such as rototilling narrow strips through beds may be helpful.

Flowering Date: May in northern Iowa

Seed maturity/Harvest date: late May to mid June in northern Iowa

Seed retention: There is a risk of seed loss from animals consuming the fruit or from fungal diseases; during the fruiting season (2-3 weeks), berries must be picked every 2-3 days.

Harvest date range at TPC: May 26 to June 13 (first harvest, 2024)

Recommended Harvest Method: Hand pick every other day. The labor required to harvest the small berries is significant: it took over 40 person-hours to gather ripe berries from an 840 sq ft plot every few days over a 19 day period, yielding 11.52 lbs of fruit and 5.47 oz of clean seed.

Appendix B

Seed cleaning and storage

Cleaning process: Place 2 parts water to 1 part fruit in a blender and process for 30 seconds. (Blender blades do not need to be altered or wrapped.) Pour the resulting mash into a 5 gallon bucket and add additional water. Stir, then allow the filled seed to settle out. Pour off the floating material, being careful to save the heavy seed at the bottom (J. Carstens, USDA-NCRPIS, personal communication, January 10, 2022). Spread the seed onto muslin cloth and place in front of a fan to dry. Airscreening will remove remaining debris.

Seed storage: cool/dry (33-50° F, 30-50% RH)

Appendix C

Released Germplasm

Source Identified material: Natural Selections/Iowa Ecotype Zone NI

The map below shows the collection locations for populations used in development of this ecotype on a base map of the Generalized Provisional Seed Transfer Zones of the US Forest Service.

A map showing 11 collection locations of Virginia strawberry on a base map of the US Forest Service Generalized Provisional Seed Transfer Zones

References

Chayka, Katy. (n.d.). Fragaria virginiana (wild strawberry). Minnesota Wildflowers. https://www.minnesotawildflowers.info/flower/wild-strawberry

Cochrane, T. S., Elliot, K., & Lipke, C. S. (2014). Wild strawberry. In Prairie plants of the University of Wisconsin-Madison Arboretum (3rd ed., p. 301). University of Wisconsin-Madison Arboretum.

Gleason, H. A., & Cronquist, A. (1991). Rosaceae. In Manual of Vascular Plants of Northeastern United States and Adjacent Canada (2nd ed., p. 242). The New York Botanical Garden.

Kartesz, J.T., The Biota of North America Program (BONAP). 2023. North American Plant Atlas. (http://bonap.net/napa). Chapel Hill, N.C. [maps generated from Kartesz, J.T. 2023. Floristic Synthesis of North America, Version 1.0. Biota of North America Program (BONAP). (in press)]

NatureServe. 2024. NatureServe Network Biodiversity Location Data accessed through NatureServe Explorer [web application]. NatureServe, Arlington, Virginia. Available https://explorer.natureserve.org/. (Accessed: February 28, 2024).

Staudt, G. (2020, July 30). Fragaria virginiana (Miller). Flora of North America. http://dev.floranorthamerica.org/Fragaria_virginiana

University of Minnesota Extension. (2025). Strawberry Farming. https://extension.umn.edu/fruit-and-vegetable-farming/strawberry-farming

USDA NRCS National Plant Data Team. (n.d.). Fragaria virginiana Duchesne. USDA plants database. https://plants.usda.gov/home/plantProfile?symbol=FRVI

Species Guide Updated 2/3/2025

common boneset

common boneset sagem Fri, 03/01/2024 - 09:44

Eupatorium perfoliatum L.

Alternate Common Names: boneset, thoroughwort

Scientific Synonyms: Eupatorium chapmanii Small, Eupatorium perfoliatum var. colpophilum Fernald & Griscom, Eupatorium perfoliatum var. cuneatum Engelmann

Family: ‌aster or sunflower family (Asteraceae)

Functional Group: forbs (wildflowers)

Printable PDF Eupatorium perfoliatum