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PLANT PROPAGATION
Laboratory Manual
Copyright © 2000, 2001, 2002, 2003, 2004,
2006
Amended Fall 2006
- All Rights Reserved -
James B Calkins, Ph.D.
Emily E. Hoover, Ph.D.
CopyrCopyt © 2000, 2001, 2002, 2003, 2004, 2006
– All Rights Reserved –
James B. Calkins, Ph.D.
Emily E. Hoover, Ph.D.
Fall Semester 2009
Department of Horticultural Science
College of Food, Agricultural, and Natural Resource
Sciences
University of Minnesota
Table of Contents
I. Introductory Information
Page
A. Plant Growth Regulators & Other Chemical Compounds Used in Plant
Propagation: General Information & Formulations ..................................................... 4
B. Sexual Propagation
1. Seed Propagation Techniques: Seed Collection, Handling, Viability, Germination,
and Germination Requirements ........................................................................................ 7
2. Propagation By Seed: Description of Seeds, Collection, Cleaning, Viability Tests,
Storage, Germination, Dormancy, Overcoming Dormancy, Environmental Factors
That Influence Germination, Seed Sowing, & Transplanting Seedlings .......................... 9
C. Asexual (Vegetative) Propagation
1. Stem Cuttings / Herbaceous Species: Herbaceous Stem Cuttings and Their
Rooting Requirements ...................................................................................................... 15
2. Stem Cuttings / Woody Species: Softwood, Semi-Hardwood, and Hardwood
Cuttings and Their Rooting Requirements...................................................................... 16
3. Leaf & Leaf-Bud Cuttings / Herbaceous & Woody Species: Leaf-Piece, Leaf-Blade,
Leaf-Petiole, and Leaf-Bud Cuttings and Their Propagation Requirements ................. 19
4. Root Cuttings / Herbaceous & Woody Species: Root Cuttings and Their Cultural
Requirements .................................................................................................................... 20
5. Vegetative Propagation Using Specialized Structures: Bulbs, Corms, Tubers,
Tuberous Roots & Stems, Rhizomes, Stolons, Runners, Layers, and Crowns .............. 21
6. Grafting and Budding: Whip & Tongue Grafts, Side Veneer Grafts, Cleft Grafts,
Approach Grafts, and Budding ........................................................................................ 23
D. Propagation Ferns and Fern Allies: Spores and Their Cultural
Requirements........................................................................................................................ 26
E. Media Effects on Propagation: Importance and Measurement of
Growing Medium Porosity and Bulk Density Characteristics .................................... 28
F. Experimental Design .......................................................................................................... 30
II.
Laboratory Exercises (E = Experiment; P = Project; T = Technique)
Page #
Individual Reports [E]
Introduction to Experimentation: Herbaceous Stem 32
Cuttings [E]
Sexual Propagation: Using Scarification to
35
Overcome Hard Seed Coat Dormancy [E]
Asexual Propagation: Adventitious Shoot Initiation
37
in Response to Treatment with BA [E]
Asexual Propagation: Propagation of Plants Using
39
Leaf Cuttings [E]
Sexual Propagation: Effects of Seed Coat Removal,
42
Treatment with GA, Seed Drying, and Cold
Stratification on the Germination of Apple Seeds
[E]
Sexual Propagation: Effects of Growing Medium on 47
Seed Germination and Seedling Performance [E]
Projects
Asexual Propagation: Propagation of Herbaceous
50
Garden and Landscape Perennials Using
Herbaceous Stem Cuttings
Asexual Propagation: Designing an Experiment to
53
Maximize Rooting of Leaf-Bud Cuttings
Sexual Propagation: Manipulation and Germination 55
of Seeds of Annual, Herbaceous Perennial, and
Woody Perennial Plants
Let’s Go Grocery Shopping: Propagation Produce
58
Techniques [T]
Sexual Propagation: Seed Morphology, Viability
61
Testing, Germination testing, and Epigeal vs.
Hypogeal Germination [T]
Asexual Propagation: Whip & Tongue Grafting [T] 64
Asexual Propagation : Unique Vegetative Structures 69
[T]
Asexual Propagation: T-Budding [T]
72
Asexual Propagation: Approach Grafting 76
Spudmatoes [T]
Asexual Propagation: Air Layering [T]
79
Starting Date
Report Due
HORT 1001 - Plant Propagation
Plant Growth Regulators & Other Chemical Compounds
Used In Plant Propagation
General Information & Formulations
The term plant hormone is very specific and often misused. Plant hormones are organic compounds,
which are naturally produced in plants (endogenous) and transported from their site of synthesis to their site
of action where they regulate physiological processes at low concentrations. Hormone biosynthesis itself
(timing, quantity, and physiological purpose including relationships to other growth substances) is
influenced by internal genetic factors and the environment. They function to regulate plant growth and
development by controlling the growth and differentiation of cells, tissues, and organs. As a group, plant
hormones are a specific subcategory of a broader classification of plant growth regulatory compounds called
plant growth regulators (PGR's). In contrast to plant hormones, plant growth regulators are applied to
plants; they do not have to be translocated, may require higher concentrations to elicit a response, and may
be either naturally or artificially synthesized. For this reason, when compounds having hormonal effects are
applied to plants exogenously they should be referred to as plant growth regulators rather than hormones.
Plant growth regulators are widely used in plant propagation and in many cases have revolutionized our
ability to successfully propagate plants by increasing the success in rooting cuttings and allowing greater
manipulation of plant tissues in micro propagation (tissue culture) systems. There are five primary types of
plant hormones and all are used as exogenously applied plant growth substances: auxins, gibberellins,
cytokinins, ethylene, and abscisic acid.
Auxins
Discovered in 1935, auxin compounds are commonly used in plant propagation to induce the production
of adventitious roots on cuttings and to control morphogenesis in micro propagation systems (tissue
culture). Along with the development of mist systems, which slow respiration and reduce water loss
(transpiration) by cooling the cuttings and increasing humidity, the discovery of auxin as a root promoter is
one of the primary advances in the ability to propagate plants from cuttings. Although a number of auxins
are synthesized by plants, indole-3-acetic acid (IAA) is the auxin most commonly produced. Auxins are
synthesized in leaf primordia, young leaves, and developing seeds and move downward in plants. They are
active in cell elongation, cell division, and cell differentiation. The stimulation of cell elongation by auxin is
involved in phototropism (growth toward light) and etiolation (stretching in the absence of light). Auxins are
also implicated in the regulation of apical dominance (inhibition of subordinate buds), cambial growth,
flower initiation and development, fruit development, and the promotion and inhibition of leaf, flower, and
fruit abscission (formation of abscission layers). Although also implicated in other processes such as the
induction of embryos in cell cultures, enhanced pollen tube growth, and tuber and bulb development, the
most important function of auxin in propagation systems is its ability to promote the initiation of
adventitious roots on cuttings.
Although it has the benefit of being soluble in water, indoleacetic acid (IAA) is unstable and breaks down
quickly when prepared and applied exogenously as a plant growth regulator. It is destroyed when exposed
to light (photooxidation) and is broken down by enzyme systems within plant tissues. For these reasons,
synthetic auxins, such as indole-3-butyric acid (IBA) and naphthalene acetic acid (NAA), which are more
stable, are more commonly used in plant propagation. Like IAA, IBA is also produced naturally in plants.
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Auxins are available in both dry and liquid formulations. Dry formulations are talc based and liquid
formulations may be water or alcohol based. In general, both are effective and easy to apply. The talc
formulations are generally cheaper; they are also easy to see on the ends of the treated cuttings. In
comparison, liquid formulations are easier to work with and tend to be more effective than talc
formulations, however, the potential for toxic effects is increased for some species when alcohol based
formulations are used.
Cytokinins
As a group, the cytokinins are believed to be involved in plant growth by stimulating cell division, cell
enlargement, and cell differentiation. In addition to kinetin, which was the first cytokinin to be identified, a
number of natural and synthetic cytokinins, including zeatin, isopentenyladenine, and benzyladenine (BA),
are known. Cytokinins appear to interact with auxins, and this interaction is widely used in plant
propagation; a high auxin/cytokinin ratio promotes rooting, a high cytokinin/auxin ratio promotes shoot
development, and high levels of both at the same time results in callus development. Cytokinins are also
thought to slow plant senescence, inhibit the breakdown of chlorophyll and proteins, and to help overcome
apical dominance and seed dormancy. The various responses associated with cytokinins likely involve
interactions with auxins and gibberellins.
Gibberellins
Gibberellins are believed to have many functions in plants. They are found at relatively high concentrations
in stem apices (especially in the leaf primordia), roots, fruits, and tubers and are known to promote shoot
growth through cell division and elongation. A lack of gibberellin may be involved in dwarfed plants.
Gibberellins are also thought to be involved in the development and germination (dormancy regulation) of
seeds and the promotion of flowering. Over 100 gibberellins have been identified in plants. GA3 and GA4+7
are the most common gibberellins used in horticulture.
Abscisic Acid
The role of abscisic acid (ABA) as it affects plant growth is mixed. ABA has been shown to have both
stimulatory and inhibitory effects, which seem to be concentration dependent. ABA is thought to be
involved in the dormancy of buds and seeds as well as plant responses to stress (especially drought). ABA is
also thought to be involved in stomatal regulation and water and ion uptake by roots. It may also influence
leaf and fruit abscission, but may not play the primary role.
Ethylene
Unlike the four groups of compounds previously discussed, ethylene is a gas under standard conditions. It
is thought to be involved in a variety of plant responses including epinasty, leaf and fruit senescence and
abscission, promotion of flowering, and overcoming apical dominance (stimulation of lateral buds).
Horticulturally, it is used to promote fruit ripening, defoliation, fruit thinning, and flowering. Ethylene has
also been used to promote the initiation of adventitious roots and to overcome seed and bud dormancy.
We will be using a variety of plant growth substances and other compounds in class. Formulations
and recipes for the preparations most commonly used follow:
5
IBA (indole butyric acid) will be available as a root promoter in both dry (talc or powder) and liquid)
formulations. IBA will be available in a commercially prepared talc formulation at concentrations of 1000,
3000, 8000, and 16,000 ppm (Hormex 1, 3, 8, & 16, respectively). IBA will also be available as a liquid
formulation in concentrations of 1000, 3000, 8000, and 16,000 ppm. For liquids, a quick-dip method of
application is used wherein cuttings are dipped for only 3 to 5 seconds. Stock solutions are stable and can
be kept indefinitely if they are kept clean; they should also be kept tightly sealed to prevent evaporation and
associated increases in concentration.
NAA (naphthalene acetic acid) will be available as a liquid formulation in concentrations of 1000, 3000,
8000, and 16,000 ppm. Although NAA is sometimes more effective than IBA in promoting root
development, it is also more likely to cause injury to the cuttings; these effects are species specific. Stock
solution characteristics, storage requirements and cutting treatment procedures are the same as for IBA.
NAD (napthylacetamide) is a synthetic auxin and will be available as a liquid formulation. NAD does not
induce as strong a reaction in plants as NAA. Although NAD can sometimes cause injury to plant material,
it tends to be more species specific than NAA. Stock solution characteristics, storage requirements, and
cutting treatment procedures are the same as for IBA.
BA (benzyladenine) will be available as a liquid formulation at a concentration of 150 and 1500 ppm.
Remember, benzyladenine is the most commonly used cytokinin in the field of plant propagation and is
used to promote the development of adventitious shoots.
GA (gibberellic acid) will be available as a potential seed treatment for overcoming dormancy in some
seeds. A commercially prepared stock solution of GA3 (Pro-Gibb) will be diluted (stock = 4% = 40,000
ppm) to a concentration of 400 ppm for use in class. Higher concentrations may also be used.
WRS (willow rooting substance) is simply a water-based extract of willow stems. Although the mode of
action is unknown, it can enhance the rooting of cuttings and may be responsible for the ease of rooting
common for willow. As a rooting promoter, it remains unquantified and effectiveness may be variable from
batch to batch. The extract is stable for long periods if refrigerated. When using WRS, cuttings are
generally allowed to take-up the extract for some time prior to sticking.
TTC (2,3,5-triphenyl-2H-tetrazolium chloride), simply referred to as tetrazolium chloride, is often used to
test the viability of seeds. When metabolized by living tissue, this colorless material is converted to an
insoluble, red compound (formazan). When applied to seeds and other plant tissues, this color change
indicates the presence of living tissue and is used as an indicator of viability. Tetrazolium chloride solutions
are prepared by dissolving TTC in water. A 1% (10,000 ppm) solution is commonly used. Seeds are
soaked for 2 to 24 hours.
H2SO4 (sulfuric acid) may be used as a pre-germination seed treatment to scarify seeds having hard,
impermeable seed coats and thereby promote germination.
Note: ppm = parts per million; a commonly used unit of measure for concentration. It is the same as milligrams per liter (mg/l); in
other words, 1ppm = 1mg/l. Also, percent (%) multiplied by 10,000 = ppm.
Always keep petri dishes containing liquid formulations of plant growth regulators covered when not in immediate use to prevent
evaporation and subsequent increases in the concentration of the growth regulator.
6
HORT 1001 - Plant Propagation
Seed Propagation Techniques:
Seed Collection, Handling, Viability, Germination Testing &
Germination Requirements
Seeds, reproductive organs composed of a protective covering, storage tissue, and an embryo, are the most
common form of reproduction for plants in the wild and a common method of propagation for cultivated
plants. In most, but not all, cases, the propagation of plants from seeds is by definition sexual propagation
since the formation of most seeds involves the transfer of pollen from the male parent to the female parent
(pollination) and the subsequent union of male and female gametes (fertilization) to form a single cell called
a zygote. Fertilization occurs within structures called ovules located within the female ovary. The zygote
begins to divide and eventually develops into an embryo (embryogenesis). As the embryo develops, stored
food reserves accumulate within the developing seed leaf or leaves (cotyledons) or separately in the form of
a tissue called endosperm. The developing seedling utilizes the stored reserves during germination and
establishment. At the same time, the ovary, which may contain multiple ovules, develops into what is
botanically called the fruit. For some plants, seeds can develop in the absence of fertilization from cells of
the female parent; when this occurs, the process is called apomixis; resulting seedlings are genetically
identical to the mother plant.
In general, producing plants from seed is relatively cheap compared to other methods of propagation. This
is perhaps the greatest advantage of propagating plants from seed. Seeds also provide a convenient and
effective way to store plants for a long period of time. Many seeds can be stored so they are on hand when
needed such as during times of the year when seed production and availability is limited. Depending on
your objectives as a propagator, there are other advantages and disadvantages associated with the
propagation of plants from seed.
One very important attribute associated with the propagation of plants from seed is the genetic diversity, or
variability, within seedling populations. This genetic diversity is the cornerstone of natural selection in the
wild and the improvement (increased vigor, higher yield, increased nutritional value, greater environmental
adaptability, enhanced pest resistance, etc.) of domesticated plants for human use through plant breeding
and selection. From a grower’s standpoint, genetic diversity and, therefore, plant variability may be looked
upon as being both desirable and disadvantageous depending on perspective. From a production
standpoint, uniformity is desirable as it simplifies the mechanics of production. If, however, the plants
being produced are destined for native plantings, native genetic diversity is preferable regarding
performance and survival.
Whether you are collecting seed yourself or purchasing seed from a supplier, a reliable source of high
quality seed is an important factor when propagating plants from seed. So, too, is the handling of the seed
following its collection or procurement and during the period of germination. When purchasing seed,
knowing the geographic location (latitude, longitude, and elevation) where the seed was collected or
produced is important. The geographic locality from which the seed was obtained is called provenance. If
available, additional information about the growing environment (soil type, pH, light, moisture conditions,
etc.) is also of interest. This information will tell you something about the potential for the resulting plants
to survive in a given environment. Knowing the source and understanding the handling and germination
requirements of specific types of seed are critical to propagation success and the long-term performance of
the plants produced.
7
Seed germination is a remarkable and complicated process. You will find that seeds differ greatly when it
comes to their specific germination requirements. Some seeds will germinate immediately without special
treatment while others may require manipulation and/or specific environmental conditions for germination
to occur. For example, seeds of most annuals, including vegetable seeds, will germinate quickly when
placed in a favorable environment (moisture, temperature, aeration, and sometimes light). Such seeds are
said to be quiescent. Seeds that are viable, but fail to germinate when exposed to conditions favorable for
germination are said to be dormant. The seeds of many woody plants and some herbaceous perennials
exhibit some type of dormancy. Seed dormancy may be caused by a variety of factors and is an
evolutionary characteristic important in the survival of many plant species. Overcoming dormancy, which
may be physically and/or physiologically induced, is often a challenge when propagating plants from seed.
Pre-germination treatments, such as scarification (breaking seed coats) and stratification (cool, moist
conditions), are often critical to success when propagating plants from seed. In addition, providing
conditions that are optimal for the germination of specific seeds can often determine success. Research
and/or hands-on experience are important in determining the proper handling and germination
requirements of specific species. Even so, germinating some seeds can remain a significant challenge even
for the experienced propagator.
Some additional things to ponder and watch for include:
 When does germination begin?
 How long does it take for all of the seeds planted to germinate (at least those that appear likely to do so)?
 When a seed germinates, what emerges first – the shoot or root?
 Is germination epigeous/epigeal or hypogeus/hypogeal?
 Is the plant to which the seed belongs a monocot (monocotyledenous) or a dicot (dicotyledenous)?
 How does the final germination percentage compare with the viability test results?
 What might be some reasons why certain seeds or groups of seeds did not germinate?
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Propagation by Seed
Description of Seeds, Collection, Cleaning, Viability Tests, Storage, Germination,
Dormancy, Overcoming Dormancy, Environmental Factors That Influence
Germination, Seed Sowing, & Transplanting Seedlings
- Excerpted from Landscape Plant Propagation Workbook: Unit III. Propagation by Seed
Dewayne L. Ingram
Sexual (seed) propagation is an important means of reproducing crop plants, including landscape plants.
Many seeds can usually be harvested from a plant; thus, many plants can be propagated from a single
mature plant. An orchid seedpod can contain over 3 million seeds but 500,000 to 1 million seeds per pod is
common in many plant species. Annuals and biennials are most commonly propagated from seed.
Sometimes seedage is the only practical means of propagating a plant, because vegetative means have not
been explored or have been unsuccessful.
Seed propagation offers genetic variability; thus, the offspring may not have the exact characteristics of the
parent plant. This allows for the selection of plants with unique features and systematic breeding for
identified characteristics. Genetic variability is a disadvantage if the goal of the producer is to grow a uniform
crop. Seedling variation is quite high in some plants, while other plants are more true to type. Techniques
using genetically stable lines or line hybrids are used to produce many of the flowering annuals and
vegetables with minimal variability.
Growers find that it takes longer to produce some plants from seed than from cuttings. Seedlings may
remain in a juvenile stage of growth for a longer period of time. Juvenile growth does not produce flowers
or fruit and such growth often has a different appearance than mature growth.
Description of Seeds: A seed is composed of three basic parts: the embryo, food storage tissue and the
seed covering. The embryo is a new plant resulting from union of male and female gamete during
fertilization. Cotyledons, or seed leaves, are attached to the embryo. Seeds are classified by the number of
cotyledons. Monocotyledons (monocots) such as grasses and palms have one cotyledon, while dicotyledons
(dicots) such as bean and peach have two. Plants such as pines are gymnosperms and their seeds may
contain as many as 15 cotyledons. A mature viable seed contains enough stored food or energy source for
seed germination and early seedling growth. The cotyledons of dicots usually contain this food reserve,
while some seeds have a mass of food reserve called endosperm surrounding or in contact with the embryo
(e.g., corn). Seed coverings can consist of the seed coat and parts of the fruit or seedpod. These structures
protect the embryo and food reserve inside the seed. They can also inhibit germination until conditions are
suitable for germination and seedling development.
Seed Collection
Seeds should be collected when ripe and just before they fall to the ground. When seeds are on the ground,
the likelihood of disease and insect infestation is greatly increased. Some seeds loose viability (ability to
germinate) soon after they are ripe and should not be collected from the ground. Several of the tropical
plants have seeds with short periods of viability. Generally, palm seeds fit into this category.
Seed maturity is often difficult to determine. Generally there is a color change of the fruit from a green to an
9
orange, black, purple or red, and the fruit may become soft. Winged seed, seedpods or fruit without fleshy
pulp may turn a brownish color as they mature and become drier. This water loss or drying coincides with
seed maturity and is a state in which the seed may be stored or distributed by wind or water.
Seed Cleaning
The pulp from a fleshy fruit may contain germination inhibitors and should be removed before seeds are
sown. The seed would probably germinate after the fruit decomposes, but the propagator may not want to
wait several weeks and the process may reduce germination. Seeds are usually removed from dried capsules
before treated or germinated.
Seeds may be cleaned individually or in bulk. Seeds may be harvested from dried capsules, or the capsules
can be crushed, if the seed is sufficiently hard, and sown with the seeds. The flesh around a seed can be
removed by hand or by some mechanical means. A procedure for cleaning several seeds at one time has
been developed by Dr. Bijan Dehgan in the Department of Environmental Horticulture at the University of
Florida. The procedure was developed for cleaning cycad seeds but will work with many seeds. The general
procedure is as follows: The fleshy coat is removed by using a long-stemmed, circular wire brush attached to
a variable speed drill. First, seeds should be soaked in water for 24 hours. Next, place seeds in large widemouth jar or a large coffee can with a plastic top in place, through which the stem of the wire brush is
passed. Add a small amount of 20-mesh sand to the seeds and enough water to cover the sand and seeds.
Turn on the drill at a slow speed and gradually increase the speed until all seed coats are removed. Wash
the seeds thoroughly. A lot of 300 to 400 seeds can be cleaned in approximately 10 to 15 minutes.
Seed Viability Tests
It is often difficult to determine from the appearance of a seed if it is alive and would germinate under the
proper conditions. There are methods of determining seed viability or potential for seed germination.
Stains can be used to determine if the seed embryo is alive. Living tissue respires in order to maintain its
integrity and tetrazolium chloride turns red when it comes into contact with respiring tissue. The seed is cut
open and the stain applied to the exposed embryo. If the embryo is alive, it will turn reddish in color.
Viable and dead seeds in some plant species can be separated by floating the seed in water. When the seed
is filled with the embryo and endosperm to support germination and early seedling growth, the seed will
sink in water. When a seed is empty or contains a relatively large amount of air, the seed will float. This is a
good means of identifying dead oak and pindo palm seeds, but it does not work with Zamia floridana,
Cycas cercinalis, coconut, fish poison tree and several other plants.
Seed Storage
Seeds of many plants can be stored and germinated weeks, months or years later. Factors affecting the
viability of seeds during storage include (1) the inherent longevity of the plant species, (2) seed moisture
content, (3) the temperature and (4) the relative humidity. The length of time seeds can be stored differs
with the plant species. Magnolia, wax myrtle and elms have short-lived seeds and may be stored only for a
few weeks or maybe up to a year without losing viability. Koelreuteria, Calendula, Petunia and Zinnia are
examples of seeds that can be stored for 2 to 15 years, and Acacia and Elaeagnus can be stored for 15 to 20
years. Research is being conducted with freezing techniques that could increase the length of time seeds can
be stored.
The moisture content of the seed should be reduced to 20 to 30 percent of seed weight before storage. This
can be accomplished by placing them in the sun or in a drying oven. Obviously, if the seed becomes too
10
dry, the embryo will die.
Many seeds will store best if a temperature 35 to 45F is provided. However, some of the tropical plants will
not tolerate temperatures below 40F. Generally, seeds from tropical plants do not store well.
The relative humidity in the storage container should be 20 to 30 percent. The moisture in the air and the
seed will become equal over time in the container; therefore, the environment in the container should be
relatively dry and the seeds should be dried somewhat before storage. Plastic bags or bottles work well as
storage containers.
Seed Germination
Although the specific characteristics of seeds differ with plant species, the general germination process is the
same. The germination process can be described in three phases: activation, digestion and cell division, and
elongation. The process begins with the imbibition or uptake of water. This may take a few hours or several
days, and the seeds will usually swell or enlarge due to the increased volume. Water uptake triggers the
activation of certain biochemical processes that result in the synthesis of the building blocks (nucleic acids,
amino acids, enzymes, etc.) for growth and development. The endosperm is digested by enzymes to
produce energy-rich compounds and nutrients that are transported to the embryo. Cell division and
elongation occurs and the developing embryo expands. The radical or root emerges from the seed by
breaking through the seed coat.
Dormancies
Many seeds are ready to germinate when the fruit is ripe or the capsule is dried. Seeds from tropical plants
usually are not dormant when mature, but many of the plants from temperate climates are dormant when
collected. Dormancies are protective mechanisms that have evolved to allow a seed to germinate at the
appropriate time. For example, dogwood seed mature in late fall but are dormant at that time. These seeds
require a cold period to satisfy some factor inside each seed before it will germinate. If dogwood seeds were
germinated in the fall, the tender seedling would probably not survive the winter. Therefore, the cold
requirement facilitates natural germination in the spring at the beginning of the growing season.
Seed dormancy can be caused by a hard seed coat that is impermeable to water or gases, or resistant to
embryo expansion. Zamia, camellia and redbud are examples of seed dormancy due to an impermeable
seed coat. Seeds from many of the stone fruits, such as peaches, are dormant due to physical restriction of
seed expansion.
The fleshy pulp, the endosperm or the seed coat can contain chemical inhibitors of germination. Some of
these inhibitors are water-soluble and can be leached out of the seed, but many cannot be washed out and
must be broken down by some chemical process. Okra seeds contain a water-soluble inhibitor that can be
leached by soaking the seeds in water for 12 to 24 hours.
The embryo may be immature when the fruit is ripe or the capsule is dry. A period of time in the proper
environment is required before germination is possible. The embryo will often increase in size during this
period and will develop the properties necessary for germination. Orchid seeds are dormant when released
by the plant, due to an immature embryo.
Some seeds have double or multiple dormancy: more than one type of dormancy. An immature embryo
may prevent germination of a seed, even though a dormancy due to a hard seed coat has been overcome. A
seed may require a cold period to break a dormancy due to the seed coat and then require a warm period
11
for development of an immature embryo. Nandina seeds require a cold-warm-cold sequence of
temperatures before germination.
Even though some seeds will germinate without a preconditioning period, the germination rate and
uniformity might be enhanced by some treatment. This condition is referred to as a physiologically
intermediate dormancy.
Overcoming Seed Dormancies
Scarification and stratification are the two most common means of breaking seed dormancy, but hot water
soaks and plant growth regulator treatments are also used. Scarification involves the breaking of the
impermeable or hard seed coat. This may be accomplished by mechanical means or by acid treatments.
Scratching the seed coat with sand paper and cutting the seed coat with a knife are suitable for some seeds.
A hammer or pliers can be used to break the seed coat in some cases. Care must be taken in any
scarification procedure not to injure the seed embryo. If a specific treatment for a particular plant species is
not known, sample seed lots treated with various techniques followed by germination tests is recommended.
Acid scarification has been used successfully in some plants to break the seed coat. Such scarification occurs
in nature when the seed coat is partially digested by fungi or bacteria or enzymes in the digestive tract of
birds and animals. Sulfuric acid is usually used for scarification. Seeds may be soaked for 5 to 60 minutes,
depending upon the toughness of the seed coat and the sensitivity of the embryo. Dry, clean seeds are
placed in a nonmetal, noncorrosive, acid-resistant container, and the acid is added slowly in a ratio of one
part seed to two parts acid. The mixture should be gently stirred intermittently during the soak to ensure
uniform results. Seeds soaked in acid should be washed thoroughly under running water for 10 minutes to
dilute the acid in contact or inside the seeds before they are sown.
Acids are extremely dangerous and proper handling procedures must be followed. Never pour water into
acids, because tremendous heat can be generated in a violent reaction. Dilute acids by pouring the acid into
water very slowly. Acid-resistant clothing including gloves should be worn when handling acids. Acids
should be disposed of properly as hazardous material and should never be poured down the sink.
Stratification is the technique of providing a moist chilling treatment to seeds to overcome a dormancy. This
technique has been used to overcome dormancies due to an impermeable seed coat or chemical inhibition.
Stratification could reduce the concentration of germination inhibitors or increase the concentration of
growth-promoting hormones. Seeds should be soaked for 12 to 24 hours and then placed in a moist
medium. Coarse vermiculite, sphagnum peat moss, equal volumes of peat and perlite, and coarse sand have
proven to be good stratification media that retain moisture yet allow aeration. Seeds should be mixed with
one to three times their volume of medium and stored in a container that provides a barrier to moisture
loss, such as a polyethylene bag. Seeds can be stratified naturally by placing them outdoors in an area or
container protected from rodents, but the temperatures outdoors may not be uniformly low enough in
Florida to provide optimum results.
Seeds are usually stratified at 32 to 40 F (0 to 2 C) for a period of 1 to 4 months. Seeds are then separated
from the stratification medium and germinated at 70 to 80 F (17 to 22 C). Germination at higher
temperatures (>90 F; >27 C) may induce a secondary dormancy.
Soaking in water will often soften seed coats and leach water-soluble inhibitors from the seed to reduce
germination time. Best results are obtained when hot water is used, but the temperature sensitivity of the
embryo differs with species. Generally, water at 170 to 212 F (67 to 88 C) is poured over seeds in a ratio
of one part seed to four or five parts water and allowed to cover the seed and cool for 12 to 24 hours.
Changing the water periodically during prolonged soaking is imperative. Seeds should not be allowed to dry
12
after the treatment but should be sown immediately.
Environmental Factors
An ample supply of high quality water must be applied during seed germination and seedling development.
There must be a proper balance between water and air in the propagation medium. Waterlogged media
cannot supply the oxygen necessary for germination and seedling growth. Monitoring the moisture level in
the medium is important. Water with excessive dissolved salts can result in poor seed germination and
seedlings growth. Seeds and seedlings differ in their sensitivity to salt levels, but generally the seedlings are
less tolerant than mature plants.
The maximum, minimum and optimum temperatures for seed germination and seedling growth differ with
plant species and maybe even cultivars within species. If the optimum temperature for a seed is not known,
the range of 75 to 80 F would be optimum for many plants and should be considered. Keep in mind that
plants native to warmer climates or those that flourish during the warmer months may have higher optimum
temperatures than plants that flourish in cooler temperatures. Reducing night temperatures 5 to 10 F (3 to
5 C) has proven beneficial in germination of some plants native to temperate climates.
Seeds can be grouped according to their requirement of light for germination. Many woody plants do not
require light for seed germination, but most epiphytes such as mistletoe and strangler fig require light. Other
species have a light requirement for germination, but this can be overcome by chilling or chemical
treatments as in lettuce, tobacco and many native weed seeds. Allium, Amaranthus and Phlox are examples
of seeds whose germination is inhibited by light. Some species such as Tsuga and Betula are sensitive to day
length, but sometimes this can be overcome by temperature treatments.
Emerging roots from a seed can absorb nutrients. The presence of nutrients in low to moderate
concentrations at this time will result in more rapid seedling growth. Care should be taken to avoid
excessive dissolved salt levels, which will injure such tender root tissue. Application of soluble fertilizers on
a periodic basis is recommended over incorporation of fertilizers in the propagation medium before the
seeds are sown.
Seed Sowing
Germinating seeds in a controlled environment such as a greenhouse and in a sterile medium will result in
optimum seed germination and early seedling development. Problems with weed seeds; nematodes, insects
and diseases can be greatly reduced with such procedures. Plastic, wood or metal flats are common
containers for seed germination, although individual containers, celled flats and preformed peat pellets are
also used. Seeds may be germinated in a flat and then the most vigorous seedlings can be transplanted to a
larger production container, or, under certain circumstances, the seeds may be sown in the production
container in which they will be sold.
The germination medium should be well drained but should hold sufficient moisture to maintain optimum
moisture in the seed. The particle size of the propagation medium components is important, because this
primarily determines moisture and aeration characteristics of the medium. The particle size must also be
considered in relation to the size of the seed to be sown in the medium. There must be sufficient contact
between the seed and the particle for exchange of moisture. A large seed can be germinated in a medium
with relatively large particles, but a small seed would settle toward the bottom of such a medium. A small
seed should be germinated in a medium with relatively small particles to provide an appropriate contact
between the seed and the particle.
The proper planting depth differs with seed. Seeds that require light for germination obviously cannot be
13
planted deeply or may not be covered at all. Generally, seed should not be planted deeper than two or
three times their diameter. Many large seed, especially tropical species like palms, may only be inserted into
the medium surface where they will remain moist.
Transplanting Seedlings
Seedlings should be transplanted before they over-grow a container and their growth habit or form is
altered. Seedlings produced in flats at a high density obviously must be transplanted earlier than seedlings
produced in larger containers at a low density. Seedlings must be hardened before transplanting. Hardening
refers to a gradual change in the environment so the seedlings can adapt to withstand more stressful
conditions than those provided in the propagation phase. The irrigation frequency is usually reduced and
the light level and fertilization may be increased. Proper hardening will ensure that the seedlings are
established in the production environment at an optimum rate.
This document is excerpted from Circular 725, Florida Cooperative Extension Service, Institute of
Food and Agricultural Sciences, University of Florida. Reviewed: April 1993. Dewayne L. Ingram,
former Professor, Environmental Horticulture, Cooperative Extension Service, Institute of Food
and Agricultural Sciences, University of Florida, Gainesville FL 32611.
Copyright Information: This document is copyrighted by the University of Florida, Institute of
Food and Agricultural Sciences (UF/IFAS) for the people of the State of Florida. UF/IFAS retains
all rights under all conventions, but permits free reproduction by all agents and offices of the
Cooperative Extension Service and the people of the State of Florida. Permission is granted to
others to use these materials in part or in full for educational purposes, provided that full credit is
given to the UF/IFAS, citing the publication, its source, and date of publication.
14
HORT 1001 - Plant Propagation
Stem Cuttings / Herbaceous Species:
Herbaceous Stem Cuttings & Their Rooting Requirements
Herbaceous stem cuttings are commonly used to propagate a wide variety of succulent, non-woody
(herbaceous) plants. In general, they are easy to root. Many species of house plants, bedding plants, and
native and introduced herbaceous landscape perennials are easily propagated using herbaceous stem
cuttings. Examples of species that are commonly propagated from herbaceous stem cuttings include
geranium (Pelargonium spp.), begonia (Begonia spp.), chrysanthemum (Dendranthema  grandiflora),
garden phlox (Phlox paniculata), dieffenbachia (Dieffenbachia spp.), and dracaena (Dracaena spp.).
As always, when preparing herbaceous stem cuttings, care should be taken to select healthy, wellconditioned plants. Stock plants are often cut back to force increased numbers of new shoots to be used as
cuttings. Cuttings should be kept cool and prevented from drying during transit and prior to sticking.
Herbaceous stem cuttings are typically 5-15 cm (2-6") long and the leaves are typically removed from the
lower portion of the cutting prior to sticking. Cuttings are typically collected by making a cut immediately
subtending (below) a node. While treatment with auxins is generally not required for rooting and usually
does not increase rooting percentages for herbaceous stem cuttings, they can improve rooting uniformity for
some species. During rooting the cuttings are maintained under humid conditions (mist or fog) to reduce
transpiration and prevent drying until the cuttings are well rooted. Cuttings from some herbaceous species
will, however, root under less controlled conditions; e.g., remember Grandma rooting cuttings in water in
the kitchen window? Most herbaceous stem cuttings are collected from terminal shoots, but cuttings may
be collected from both terminal and lower portions of the stems. Flowers that are present or appear later
should be removed so that all energy (photosynthate) is exclusively utilized for the initiation and production
of roots.
Polarity is another consideration when preparing herbaceous stem cuttings or, for that matter, any type of
cutting. Polarity has to do with maintaining the orientation of the cutting relative to its orientation on the
source plant; in other words, up vs. down (distal vs. proximal). Polarity is an inherent condition wherein the
cutting responds differently on one end than the other. For example, stem cuttings tend to form roots on
the basal end relative to their orientation when collected from the source plant. When cuttings are stuck
upside down, they still tend to form roots on what was the proximal end even though polarity has been
reversed.
15
HORT 1001 - Plant Propagation
Stem Cuttings / Woody Species:
Softwood, Semi-Hardwood, and Hardwood Cuttings &
Their Rooting Requirements
Softwood & Semi-Hardwood Cuttings
Softwood cuttings, also called summer softwood or greenwood cuttings, are collected during late spring and
early summer when the new growth of woody plants is still "soft" or unlignified. New wood remains "soft" for
a period of 2 to 8 weeks depending on species and environmental conditions. For softwood cuttings, the
source plants are still actively growing and shoots have not set terminal buds. Leaves near the tips of the
growing shoots are not completely expanded. Timing is important; the wood must not be too succulent or
too woody. Cuttings collected when the wood is too soft will often rot while those collected too late may
root poorly or not at all. For some species, timing is very critical regarding success with softwood cuttings.
For example, lilac (Syringa spp.) cuttings must be collected and stuck within a very short time window just
prior to terminal bud set to root reliably and in sufficient percentages to be profitable. Softwood cuttings
generally root easier and quicker than semi-hardwood and hardwood cuttings, but, since the cuttings are soft
and tender, they also require more care during collection, handling, and rooting. Softwood cuttings are
widely used in the propagation of many deciduous landscape shrubs and some trees and a variety of woody
tropical and subtropical plants some of which are used as houseplants. Examples of species typically rooted
from softwood cuttings include honeysuckle (Lonicera spp.), lilac (Syringa spp.), rose (Rosa spp.), spirea
(Spiraea spp.), and viburnum (Viburnum spp.).
Semi-hardwood cuttings are collected during late summer and early fall and, owing to the presence of
leaves, are similar in appearance to softwood cuttings. As the name implies, however, the condition of the
wood (soft vs. hard or mature) falls somewhere between that of softwood and hardwood cuttings. At the
time of collection, stock plants are still in leaf although terminal buds have generally been set and the leaves
along the full length of the stems are fully expanded. The wood has not completely matured, but is no
longer soft and pliable as with softwood cuttings. Many broadleaf evergreens, and several woody tropical
and subtropical species, are propagated using semi-hardwood cuttings. Although less common, a number
of deciduous species can also be propagated from semi-hardwood cuttings. Examples of species typically
rooted from semi-hardwood cuttings include euonymus (Euonymus spp.), evergreen azaleas
(Rhododendron spp.), evergreen hollies (Ilex spp.), cliff green (Paxistima canbyi), and boxwood (Buxus
spp.).
When preparing softwood and semi-hardwood cuttings, care should be taken to select healthy, wellconditioned plants. Cuttings should be kept cool and moist during transit and prior to sticking. Since the
cuttings have leaves, special care must be taken to prevent drying out during transit and storage and the
cuttings must be rooted under humid conditions (mist or fog) to reduce transpiration and prevent drying
until the cuttings are well rooted. For some species, polyethylene tents with occasional sprinkling are
sufficient to control transpiration. Bottom heat can be helpful in the spring if temperatures are cool and
treatment with plant growth substances may be helpful in promoting rooting. Cuttings are typically collected
from terminal growth, should be 8 to 15 cm (3-6") long and should have 2 or more nodes. The basal cut is
usually made just below a node. Leaves should be stripped from the basal portion of each cutting prior to
sticking. If flowers or flower buds are present, they should be removed so that all energy (photosynthate) is
exclusively utilized for the initiation and production of roots.
Several labs will investigate the difference between softwood and semi-hardwood cuttings. The objective of
16
these labs is the production of entire plants from softwood and semi-hardwood cuttings and to compare
rooting for these two types of cuttings. By the time classes begin in the fall, we are way beyond the optimal
time to collect and stick softwood and most semi-hardwood cuttings. We will, therefore be using container
grown plants that have been maintained under conditions that will keep them from going dormant. In this
way we can provide plants with new growth suitable for use as softwood cuttings and semi-hardwood
cuttings. Observe your cuttings at least weekly and record your observations.
Hardwood Cuttings
Hardwood cuttings can be successfully used to propagate a number of woody trees, shrubs, and vines. The
wood used is completely mature or woody and the cuttings are typically collected from dormant plants in
late fall, winter, or early spring. Exactly when the cuttings are collected may influence how the cuttings are
handled as an artificial cold treatment may be needed to initiate shoot growth for cuttings collected prior to
exposure to sufficient chilling (a period of exposure to temperatures just above freezing required to
overcome dormancy). Many deciduous and narrow-leaved evergreen species and some broad-leaved
evergreens are propagated using hardwood cuttings. Examples of species typically propagated from
hardwood cuttings include honeysuckle (Lonicera spp.), spirea (Spiraea spp.), forsythia (Forsythia spp.),
grape (Vitis spp.), currants and gooseberries (Ribes spp.), juniper (Juniperus spp.), white cedar (Thuja spp.),
false cypress (Chamaecyparis spp.) and yew (Taxus spp.).
As with any propagation technique, when preparing hardwood cuttings care should be taken to select
healthy, well-conditioned plants. Although drying-out is of considerably less concern than with other, more
succulent, types of cuttings, hardwood cuttings are best kept cool and should be prevented from drying
during transit and prior to sticking. Cuttings that are stuck in early spring may be collected just prior to
sticking or they may be collected in the fall and stored outdoors or indoors under more controlled
conditions (cool temperatures and moist conditions to prevent drying). In areas with severe winters,
collection and storage of cuttings in the fall or early winter, prior to the onset of potentially lethal winter
temperatures is advisable. This is especially true for species that are not reliably cold hardy.
For most species native to north temperate regions, exposure to a period of cool (34-38F) temperatures is
required to overcome dormancy and initiate bud break and the resumption of normal growth in the spring.
For such species, the wood used to prepare hardwood cuttings must also be exposed to such conditions.
This cold treatment occurs naturally during the winter for cuttings collected in the late winter or early spring
or is provided artificially through refrigerated storage for cuttings collected in the fall or early winter. In
some cases, prepared cuttings are stored under relatively warm conditions for a time to induce callus
formation prior to sticking to hasten rooting. Remember that depending on species and the time of
collection, such cuttings may also require a period of chilling, either before or after callusing, to overcome
dormancy. For some species, hardwood cuttings may be collected and stuck directly in outdoor fields or
beds in the fall or early spring. While the practice is used to a limited extent in northern climates such as
Minnesota, it is quite common in warmer areas such as the central and southern U.S.
Hardwood cuttings are typically rooted under humid conditions (mist or fog) to reduce transpiration and
prevent drying until the cuttings are well rooted. In some cases, shading the cuttings, humidity tents, regular
sprinkling, or a combination of these methods can provide sufficient reduction in water loss. Bottom heat,
wounding, and treatment with plant growth substances are often helpful in promoting rooting of hardwood
cuttings. The concentrations of rooting promoters used are typically higher than those used for softwood
and semi-hardwood cuttings. Cuttings may be collected from both terminal and lower potions of the stems.
Age of the stem may affect rooting success. Cuttings that include a short section of stem from older wood
are called mallet cuttings while those that include only a small piece of older wood torn from the adjacent
stem are called heel cuttings. Hardwood cuttings usually have 2 or more nodes and may be anywhere
between 10 and 76 cm (4-30") long. The basal cut is usually made just below a node. In the case of
17
hardwood cuttings, which initially are void of leaves, much of the success is dependant on the stored food
reserves, mainly carbohydrates, present in the cutting. If flower buds are present and are recognizable, they
should be removed, along with any flowers that appear following bud break, so that all energy (stored
carbohydrate and new photosynthate) is exclusively utilized for rooting.
Cutting orientation is very important with hardwood cuttings. Because the leaves are not present, make sure
to observe the buds. In all cases, the buds should be pointing upright to assure correct polarity (up vs. down
relative to the orientation of the cutting on the stock plant).
18
HORT 1001 - Plant Propagation
Leaf & Leaf-Bud Cuttings / Herbaceous &
Woody Species:
Leaf-Piece, Leaf-Blade, Leaf-Petiole, and Leaf-Bud Cuttings &
Their Propagation Requirements
Herbaceous Species: Leaf-Piece, Leaf-Blade, and Leaf-Petiole Cuttings
Some plants can be propagated using leaf cuttings. Several different types of leaf cuttings are used as
determined by the amount of leaf tissue used. They include leaf-blade cuttings, which consist of the leaf
blade only, leaf-petiole cuttings, which include the leaf blade and the attached petiole, and leaf-piece
cuttings, which are comprised of only pieces of leaves. Although somewhat species dependent, the
potential to produce many new daughter plants from a limited number of stock plants, or even a single
stock plant, is very high when leaf cuttings are used owing to the small size of the propagules involved.
Since no buds or roots are present on the cuttings, successful propagation from leaf cuttings requires the
development of both adventitious roots and shoots. The portions of the leaf used for propagation purposes
generally do not remain as a permanent part of the new plants produced. Depending on species, plant
growth substances have had mixed results on propagation success with leaf-piece, leaf-blade, and leaf-petiole
cuttings. Many species, mostly herbaceous, that are used as houseplants or bedding plants can be
propagated from leaf cuttings. Some examples of species that can be propagated from various types of leaf
cuttings include begonia (Begonia spp.), Cape primrose (Streptocarpus spp.), African violet (Saintpaulia
spp.), snake plant (Sansevieria spp.), and peperomia (Peperomia spp.).
Herbaceous & Woody Species: Leaf-Bud Cuttings
While they may root, some plants are unable to initiate adventitious shoots from leaf-piece, leaf-blade, or
leaf-petiole cuttings and the rooted cuttings, therefore, never develop into new plants. Many of these plants
can, however, be propagated easily from cuttings if the propagule includes a preformed shoot meristem.
They can, for example, typically be propagated from stem cuttings. For some of these plants, a cutting that
includes the node portion of the stem along with the attached leaf is all that is needed to be successful.
Such cuttings, consisting of the leaf blade, the petiole, the node, and the associated axillary bud, are called
leaf-bud cuttings or single-node cuttings. The bud portion of the cutting contains the new shoot primordia
and only roots need to be initiated to produce a new plant. Many species, including most herbaceous
species that are used as house or bedding plants, can be propagated from leaf-bud cuttings. Leaf-bud
cuttings are also used to commercially propagate a number of woody species. Some examples of species
that can be propagated from leaf-bud cuttings include peperomia (Peperomia spp.), geranium (Pelargonium
spp.), rhododendron (Rhododendron spp.), red maple (Acer rubrum), clematis (Clematis spp.), and
blackberry (Rubus alleghaniensis).
As always, leaf-piece, leaf-blade, leaf-petiole, and leaf-bud cuttings should only be collected from healthy,
well-conditioned plants. The cuttings should be kept cool and prevented from drying during transit and
prior to sticking. The cuttings are maintained under high humidity (usually under mist or in fog houses) to
reduce transpiration until the requisite roots and shoots are produced.
19
HORT 1001 - Plant Propagation
Root Cuttings / Herbaceous &
Woody Species:
Root Cuttings & Their Cultural Requirements
For some plants, root pieces can be used to vegetatively propagate new plants. True root cuttings must be
distinguished from rhizomes, which, although they are often mistaken as roots and can certainly be used as
propagules, are by definition modified, underground stems. The common spice we know as ginger
(Zingiber officinale) is a good example. Although it is called ginger root, it is a rhizomatous, tropical herb
and a good example of a plant that can be easily propagated from rhizomes, but not true roots. Rhizomes
have nodes and associated bud initials. Root cuttings do not have nodes and must initiate adventitious
shoots if they are to be used as a method of propagation.
When compared to most other methods of propagation, propagation using root cuttings is relatively
uncommon. The list of plants that have the capacity to initiate shoots from root tissue is limited and,
because roots are located below ground, root cuttings are more difficult to collect. For most plants, other,
more effective and economical propagation methods are available. In certain cases and under certain
circumstances, however, root cuttings are the method of choice.
Depending on species, root cuttings may be collected at various times during the year. Most are collected
during late summer and early fall. While root cuttings can also be collected during the winter, unless the
soil has been prevented from freezing by mulching heavily, frozen ground poses a problem in colder
regions. As a general rule, any plant that suckers from the roots is a likely candidate for propagation from
root cuttings. Some examples of species that can be propagated from root cuttings include sumac (Rhus
spp.), garden phlox (Phlox paniculata), devil's walking stick (Aralia spinosa), Oriental poppy (Papaver
orientale), horseradish (Armoracia rusticana), sweet fern (Comptonia peregrina), and red raspberry (Rubus
ideaus).
As with other types of cuttings, root cuttings should only be collected from healthy, well-conditioned plants.
The root cuttings should be kept cool and prevented from drying during transit and prior to sticking.
Unlike stem cuttings, where polarity (direction of growth; up vs. down) can usually be easily determined
from the orientation of the buds, when working with root cuttings it is important to indicate proper polarity
in some fashion at the time of collection. Knowing polarity is generally of greater concern for plants with
large, fleshy roots than those with thin, delicate roots. To this end, distal ends of root cuttings are typically
cut on the bias while the proximal ends are given a flush cut. Cuttings are typically 5 to 20 cm (2-8") long
and will vary greatly in thickness depending on species (actual length is often correlated with diameter;
shorter cuttings for thicker roots and vice versa). Cuttings from plants with large fleshy roots are either stuck
upright or inclined on their sides, proximal end up, in shallow trenches cut into the rooting medium. The
cuttings are then covered with medium such that their tops are just below the surface. Cuttings from plants
with thin, fibrous roots are typically planted horizontally just below the surface. The cuttings are kept moist
and maintained under optimal conditions until shoots and additional roots are produced.
20
HORT 1001 - Plant Propagation
Vegetative Propagation Using Specialized
Vegetative Structures
Bulbs, Corms, Tubers, Tuberous Roots & Stems, Rhizomes,
Stolons, Runners, Layers, and Crowns
Specialized vegetative structures such as bulbs, corms, tubers, tuberous roots and stems, and pseudobulbs
are mainly associated with vegetative propagation of herbaceous species. Rhizomes, stolons, runners, layers,
and crowns may be used to propagate both herbaceous and woody species. Some of these structures,
including bulbs, corms, tuberous roots and stems, and some rhizomes, function primarily in food storage
during dormant periods in addition to serving a reproductive function. Rhizomes (modified, underground
stems) and stolons (horizontal stems that creep across the ground) root and produce shoots at the nodes
and can serve both storage and reproductive functions. These vegetative structures can be quite short,
resulting in clumps, or distinctly elongated resulting in rather large colonies. A runner is a specialized type
of stolon, which roots and forms a new plant at its tip when it contacts the soil. In addition to serving a
storage function, bulbs (tulip, lily) and corms (crocus, gladiolus), produce naturally detachable offshoots
called offsets, which become or can be artificially separated from the parent structure (a form of division).
This natural form of reproduction can also be exploited commercially by inducing the formation of offsets,
which are then removed from the parent structure and grown on to produce new plants.
Scooping, Scoring, Chipping, Scaling, & Twin Scaling
Bulbs are produced by monocotyledonous plants and function in storage and reproduction. Their
structure is defined by a short stem terminated by a growing point or flower primordia enclosed by thick,
fleshy, modified leaves or scales. Bulbs and may be classified as being either tunicate (also referred to as
laminate) or non-tunicate (also referred to as scaly). Tunicate bulbs are protected by a layer of dry,
membranous bulb scales called the tunic. The inner scales are arranged in tight, contiguous layers or
lamina resulting in a rather solid structure. A basal plate can be found at the base of each bulb and it is
from here that new roots develop annually. Onion is perhaps the most recognizable example of a tunicate
bulb. Tulip and daffodil are also examples of tunicate bulbs. Non-tunicate, or scaly, bulbs lack the outer
layer of dry scales and the fleshy scales are clearly visible and distinctly individual. Having a less solid
structure, and lacking the tunic, scaly bulbs are more easily damaged and susceptible to drying compared to
tunicate bulbs. Roots are also formed at the base of scaly bulbs each year, but they live for two seasons.
Lilies are the best examples of scaly bulbs.
Scooping, scoring, chipping, and scaling are common methods used in the propagation of plants that
produce bulbs. These methods induce the production of offsets or, in this case, bulblets. Scooping
involves the mechanical removal of the basal plate, which exposes the bases of the bulb scales inside the
bulb. The bulbs are callused in the open or in a dry medium for a few days at 21C (70F). This reduces
the chances of infection and decay. The bulbs are then incubated at higher humidity by covering them with
a moist substrate, such as moistened peat, to encourage the development of adventitious bulblets at the
bases of the exposed bulb scales. The bulblets are then removed, planted, and grown-on to larger size.
Scoring is the practice of making cuts through the basal plate of the bulb. As with scooping, the bulbs are
allowed to callus for a few days and then incubated at higher humidity. Bulblets are initiated in the axils of
21
the bulb scales along the cuts. The process involves making three cuts, in the pattern of an asterisk, through
the basal plate using a sharp knife. Once again the bulblets are removed and grown-on to a larger size.
Scaling involves the peeling-off of individual bulb scales from the mother bulb for use in propagating new
plants. The bulb scales may be treated with a fungicide and are then planted or incubated in moist peat
moss or sand at temperatures of 18-21C (65-70F) for a period of a few weeks or more depending on
species and treatment. The individual scales will root and grow. In addition bulblets will be initiated at the
base of the scales. Most lilies can be propagated by scaling. When two or sometimes more scales are used
in propagation, by dividing the bulb into several slabs or smaller sections each with a piece of basal plate,
the process is called twin scaling or chipping, respectively. Grape hyacinth are easily propagated by twin
scaling and narcissus by chipping.
Some bulbous plants also produce small "bulbs" along their stems. When produced above ground they are
called bulbils; when produced below ground they are called bulblets. Both may be produced naturally or
induced by plant manipulation. All bulbous species naturally produce bulblets. Lily is probably the best
example of a plant capable of producing bulbils.
Layering
Layering is another method of vegetative or asexual propagation that occurs naturally or under controlled
conditions. Layering is the rooting of shoots while they are still attached to the parent plant. Layers may be
natural or induced and can be used to propagate both herbaceous and woody plants. In cultivation, the
process involves covering a portion of the stem with moist soil or some other medium in order to promote
rooting. Wounding and/or treatment with root promoters are often involved. Air layering, tip layering,
mound layering or stooling, trench layering, and serpentine layering are the most common types. Stolons,
runners, and rhizomes might be considered naturally rooted layers. Rooted suckers produced from stem
tissues are also essentially natural layers.
Division
Division is perhaps the simplest form of asexual propagation. It is simply the division of the parent or
mother plant into two or more viable sections, which are then planted. The removal of offsets from bulbs
can be considered a type of division. Other examples of plants propagated by division include plants having
tubers (caladium, potato), tuberous roots (dahlia, sweet potato), tuberous stems (tuberous begonia,
cyclamen), rhizomes (lily-of-the-valley and iris), stolons (bugleweed, club mosses), runners (strawberry,
spider plant, boston fern), or branched crowns (most herbaceous perennials and some woody shrubs). The
crowns of some plants may also produce offshoots, which are similarly divided into separate plants. Some
plants can also be propagated from naturally occurring suckers (adventitious shoots arising from below
ground) most often from root tissues (sumac, raspberry). Finally, cormaceous plants, such as gayfeather,
can also be propagated by division.
22
HORT 1001 - Plant Propagation
Grafting & Budding
Whip & Tongue Grafts, Side-Veneer Grafts, Cleft Grafts,
Approach Grafts, and Budding
Grafting / Graftage
Grafting is the art and science of combining two or more pieces of plant tissue such that they knit together
to form a complete and viable plant. Grafting takes its cue from and is an adaptation of the ability of some
plants to naturally form graft unions in the wild. Grafting is used to propagate selected landscape plants and
fruit trees that are otherwise difficult to propagate by other, more economical, means such as the use of
cuttings. Grafting is also used in cases where it is desired and useful to combine two or more genotypes to
produce superior plants. Grafting a desired top onto a specific rootstock can confer special characteristics
to the finished plant such as dwarfing, enhanced vigor, precocious flowering, enhanced mineral nutrition, or
tolerance to specific soil conditions. In addition, grafting can be used to influence plant form, as when a
weeping form is grafted onto a standard. Grafting can also be used to repair damage to existing plants that
are valuable in the landscape or a production setting. Finally, graftage can be employed in the creation of
unique plants or multiple varieties (e.g., 5 in 1 apple) which are produced mainly as curiosities.
The plant tissues most often used in grafting include a scion piece and a rootstock. The scion consists of a
portion of the crown of an existing plant; it may be as small as a single bud. The rootstock may consist of
an entire root system or simply pieces of roots depending on the type of graft and its purpose. If the graft is
successful, the scion is destined to become the growing shoot while the rootstock will serve as the resulting
plant's root system. Sometimes additional pieces of stem are inserted between the primary scion and
rootstock; they are called interstocks and serve to introduce additional characteristics (such as dwarfing) into
the finished product. Interstocks are most commonly used in the development of improved fruit trees.
It is the ability of the cambial tissues within the scion, interstock, and rootstock to produce callus tissue
which differentiates into functional xylem and phloem and leads to the development of a viable, continuous
connection between the cambial tissues of the grafted parts that enables grafting to serve as a practical means
of propagation. To this end, it is very important that the stock and scion pieces fit closely together and be
held securely in place until the graft is sufficiently healed.
In most instances, the scion is grafted onto a stock that has an existing root system. Such rootstocks may be
rooted cuttings, seedlings, transplants, or even much older, larger plants. Either the entire root system or
just parts of the root system (e.g., piece rooting) from the stock plant may become part of the finished graft.
The technique whereby a scion is grafted onto an unrooted cutting, and the cutting is rooted at the same
time as the graft heals, is called stenting. Although we will be using this technique in class, and it does have
some specific commercial uses, stenting is a relatively uncommon practice in the real world. Grafting is
labor intensive and, from a commercial perspective, success rates must be in the range of 90% or more to
be economical. Grafting is, therefore, limited to high value plants that cannot be propagated using less
expensive methods. The rooting of cuttings is also a more costly venture than growing plants from seed.
Only very high success rates or the production of a grafted plant with superior rootstock can, therefore,
justify stenting as a method of producing grafted plants. Although it is a viable technique in certain instances
(e.g., nurse grafting), these considerations explain the relatively uncommon commercial use of stenting; the
costs involved require that the rooting success for the grafted cuttings be essentially 100% since the grafting
costs are typically already limiting.
23
Compatibility between the scion and rootstock is of paramount importance in the success of any graft. In
general, the more closely related the two parts (scion & rootstock), the more likely the graft will be
successful. Therefore, grafts between different plants within a species are typically compatible, grafts made
between different species within the same genus are also frequently successful, and, although much less
common, unions between different genera within the same family are also sometimes successful. Grafts
between plants separated by more than a family relationship will almost always fail completely. The
relationship between grafting success and plant taxonomy is further illustrated by the following examples: A
specific variety or cultivar of green ash (Fraxinus pennsylvanica; Oleaceae, Olive Family) budded onto a
seedling green ash rootstock (= same species) will be successful; white ash (Fraxinus americana) grafted onto
a green ash rootstock (= same genus) will also typically be successful; and common privet (Ligustrum
vulgare; Oleaceae, Olive Family) grafted onto a green ash rootstock (= same family) will also usually be
successful. Green ash grafted onto a sugar maple (Acer saccharum; Aceraceae, Maple Family) rootstock (=
different families) will, however, prove unsuccessful.
In addition to compatibility between the species to be grafted, the inherent ability of the individual species
being grafted to produce callus will also influence grafting success. Other factors, including the selection of
healthy stock plants, relatively warm temperatures, good aeration, and adequate moisture (high humidity)
are all required for grafting success. Except for the expertise of the grafter in preparing the stock and scion,
these factors are most often responsible for the success or failure of grafted plants.
Budding
Budding is a specialized form of grafting wherein the scion has been reduced to a single bud from the plant
to be propagated. The technique is widely used in the production of landscape plants; mainly named tree
cultivars (e.g., Acer platanoides `Deborah', Malus `Spring Snow'). Budding is also commonly used as a
method of propagating various fruit cultivars. In most cases, the single-bud scion is typically budded onto a
seedling rootstock grown specifically for that purpose. The stems that provide the scion buds are called bud
sticks or bud wood. In most cases, the bud is grafted onto the stock near the soil line and the top of the
rootstock is removed once the grafted bud begins to grow. Reducing the size of the scion to a single bud
greatly increases the efficiency of propagating the parent plant. Although roses are also commonly budded,
the practice is becoming less common in favor of cuttings and the subsequent production of plants on their
own roots.
We will investigate two budding techniques in class – T-budding (also called shield budding) and chip
budding. The main difference between the two is the condition of the understock, which is essentially
determined by the time of year that the budding operation is performed. T-budding is done when the bark
is said to be slipping - a time when cambial growth is active and the bark is, therefore, easily separated from
the underlying wood. Chip-budding is used when these conditions are lacking and the bark is not easily
separated from the wood. Although there are other forms of budding, T-budding and chip-budding are the
primary budding techniques used.
Cleft Grafting
An infrequently used, but at times very valuable type of graft, mostly used in fruit production is cleft grafting.
Cleft grafting is often referred to as ‘top—working’. Sometimes growers are faced with a recently planted
block of trees of a cultivar that is under performing. As opposed to pushing the whole block out and
replanting, some growers will opt to use the established planting to support the growth of new more
desirable cultivar. By ‘top-working’ the existing trees they save the price of removing the existing orchard,
purchasing and planting new trees. In addition, they save considerable time in getting the new cultivars into
production. We will be cleft grafting apple scion wood onto simulated apple rootstocks.
24
Spudmatoes?!
Although relatively uncommon in practice, many herbaceous species can also be grafted. As with woody
species, various types of grafts can be used and the more closely related the stock and scion the more likely
a graft will be successful. For example, various types of squash, say acorn squash, or butternut squash, or
pumpkin (all Cucurbita spp.; Cucurbitaceae, Gourd Family), can be grafted to one another successfully.
Such grafts can be used to create novelty plant combinations, impart disease resistance, or to improve the
ability of plants to grow under specific conditions.
25
HORT 1001 - Plant Propagation
Propagating Ferns & Fern Allies:
Spores & Their Cultural Requirements
Ferns are unique and often treasured plants both in the wild and in cultivated landscape and garden settings.
All true ferns were at one time included in the same family, the Polypodiaceae, but have since been
separated into several families by some recent authorities. Family classifications are, therefore and
unfortunately, somewhat confused in the literature. Most references still list most true ferns as members of
the Family Polypodiaceae. The club mosses (Lycopodiaceae; also called ground pines and running pines
or cedars) and horsetails (Equisetaceae), also common in the wild and sometimes planted in the landscape,
are often grouped together with ferns and referred to as the fern allies. Although perhaps not as closely
related as once thought, they are also non-flowering plants and have life cycles that are similar to that
observed for ferns.
Classified as non-flowering plants, ferns and their allies do not produce flowers or seeds. They are,
however, classified as vascular plants owing to the presence of conducting tissues for the transport of water
and minerals (xylem) and photosynthates (phloem). As a group, these interesting plants typically posses
true roots and leaves (in the case of ferns, fronds) both of which originate from a perennial rhizome. The
rhizome may grow horizontally or vertically and may be found below or above ground depending on
species. The “leaf”, or frond, is divided into two main parts – the stipe (the leaf stalk or petiole) and the
blade (the leafy, expanded portion of the frond). In addition, the portion of the stipe to which the leaflets
are attached is sometimes referred to as the axis or rachis. Depending on species, the blade can be quite
variable; it may be simply lobed or divided into smaller sections (or leaflets) called pinnae, pinnules, or
pinnulets depending on the degree of division.
The life cycle of ferns and other non-flowering plants makes them unique, but has served well for hundreds
of millions of years. In the wild, reproduction occurs by both vegetative and sexual means. The complete
life cycle of the ferns and fern allies is characterized by two, independent, distinctly different, alternating
generations of plants, one of which produces sex cells while the other produces spores. The familiar, sporebearing plants we call ferns are examples of the sporophytic or spore-producing generation. The
sporophyte can also propagate itself vegetatively by means of its rhizome. Spores are comprised of a single
cell containing nucleated protoplasm and are produced in miniature, stalked (sporangiophore) spore sacs
called sporangia, which are produced on either the underside of the fronds, or on separate spore-bearing
fronds. Fronds that bear spores are called fertile fronds or sporophylls. When produced on the
undersides of the leaves, the sporangia are typically clustered in covered structures called sori. Each sorus
looks like a small dot and as a group the sori are typically arranged in an organized pattern on the
undersides of mature fronds. Millions of spores are produced, but few will land in a spot suitable for
growth. Under favorable conditions, spores germinate to produce small, free-living plants called prothallia,
which comprise the sexual generation and look nothing like what we think of when we think of ferns. Each
prothallium (also prothallus), or gametophyte, consists of a small, inconspicuous, green, heart-shaped
structure and grows flat on the soil surface. Separate male and female sex organs (antheridia and
archegonia, respectively), which produce eggs and sperm (also called spermatozoids or antherozoids), are
produced on the underside of the prothallium. In the presence of moisture, the motile sperm swim about
and unite with an egg produced by the archegonium. The resulting zygote then begins to divide and
develops into the familiar spore-bearing plant (sporophyte) we call a fern. The entire process is relatively
slow and generally requires a period of several years.
In cultivation, ferns may be propagated by a variety of means depending on the goals of the propagator and
26
the species of fern involved. Division of clumps or the rhizome is the most common for plants grown in the
garden and where plants that are identical to the parents are desired. When large numbers of plants are
desired and genetic variability is of less concern or desired, spores are used. Some ferns naturally produce
or ban be stimulated to produce bulbils in the axils of their leaves which can be used to propagate new
plants. As with plants propagated by division, the plants produced are genetically identical to the parent.
Time is also of concern; division will produce full size plants the quickest, followed by bulbils, followed by
spores, which will require several years.
The first step in propagating ferns from spores is the collection of the spores. When the spores are ripe
and ready to be released, fertile fronds are collected and placed in a clean dry container. They are allowed
to dry for several days and then lightly tapped or shaken to dislodge the spores. Fern spores are often
viable for only a short period of time (days) and should, therefore, be sown as soon after harvest as possible.
If immediate sowing is not possible, most spores can be stored and viability maintained under refrigeration.
Sowing is accomplished by filling a small container with moist, sterile, growing medium and carefully
sprinkling the spores onto the surface of the medium. The culture must be kept in a moist, sterile
environment in indirect light. Temperatures of 60 to 70ºF (15 to 20ºC) are recommended. Following
germination of the spores, the developing prothalli will appear as green film on the surface of the medium.
When small fronds appear, the young ferns are transplanted and gradually hardened off.
The American fern Society is an excellent source of information about ferns. Visit their web page for more
information about ferns (http://www.amerfernsoc.org).
27
HORT 1001
Plant Propagation
Media Effects on Propagation:
Importance & Measurement of Growing Medium
Porosity & Bulk Density Characteristics
A carefully planned growing medium, specifically suited to the plants being grown, is one of the critical
aspects of successful plant propagation and production in general. The characteristics of a growing medium
become even more critical for plants grown in containers where the volume of the container limits the
growing environment on many levels. Special care must, therefore, be focused on all aspects of production
to optimize plant growth and performance in containers. Media porosity and bulk density are just two of
the many factors that must be considered when selecting a growing medium.
Porosity
Growing medium porosity characteristics are but a single, yet highly important, group of production factors
that influence plant performance or may even determine the success or failure of an entire crop. When
choosing a growing medium our goal is to strike a balance between aeration and moisture holding capacity
such that the medium provides sufficient oxygen and moisture for plant roots without requiring excessively
frequent moisture replenishment (rainfall or irrigation). It should be noted that additional considerations
such as fertility, pH, and weight, are also involved in designing a proper growing medium.
The total porosity of a container medium is a measure of the space within the container that is not occupied
by the solid medium components under standard moisture and planting or growing conditions. In other
words it is the space within the medium that is occupied by either water (water-retention porosity) or air
(aeration porosity) at the time of measurement. Porosity measurements are standardized such that they are
measured for a growing medium at field or, in this case, container capacity. Field or container capacity is
when the medium contains as much water as it can possibly hold just after it has been fully saturated and
then drained.
Desired Values:
Media porosity is a dynamic characteristic; it is influenced by the characteristics of the growing medium and
container depth. The porosity characteristics of a growing medium also change over time in response to
settling and the degradation of organic components. As a growing medium ages in a container, aeration
porosity typically decreases while water retention porosity increases. The porosity characteristics of a given
medium are directly related to the depth of the containers in which it is used; a growing medium will always
hold less air and more water in a shallow container compared to a deeper container of the same volume.
When it comes to plant growth and performance, porosity requirements are also species specific since
plants differ in the capacity of their roots to tolerate low oxygen, high moisture conditions.
28
In general:
Aeration Porosity (should be at least 10%) – 15-30% (perhaps higher; up to 45%)
Water-Retention Porosity – 35-50%
Total Porosity – 50-80% (sometimes up to 95%)
Bulk Density
Bulk density is the mass or weight per unit volume (in situ) of oven-dry growing medium and is commonly
expressed as g/cm3. It is an important characteristic of any growing medium whether it be soil or artificial.
Bulk density is determined by a variety of factors including the type and size of the particles that make up
the medium, how tightly the particles fit or are packed together, as well as organic matter content, moisture
content, and biological activity which influences the aggregation of particles and ultimately determines the
structure of the growing medium (how the particles are held together and arranged). These characteristics
in turn influence medium porosity and subsequently aeration and soil moisture characteristics. Bulk density
also influences water infiltration and thereby mediates erosion potential and soil moisture water recharge.
The relationships between bulk density and porosity, and their effects on plant performance, are complex.
Typical & Desired Values:
The bulk density of a growing medium is dependent on the texture of the various medium
components and the structure of the medium (i.e., how the medium components are organized).
Bulk density is also dependent on the degree of compaction and the effects of soil moisture on
shrinkage and contraction of the growing medium. The effects of bulk density on plant
performance are also species specific as plants differ in their capacity to tolerate compacted soils.
A bulk density of:
0.1 to 0.6 - is typical of organic soils
1.0 to 1.3 - is typical of granulated clay surface soils
1.3 to 1.8 - is typical of course textured (sandy) surface soils
In general:
A bulk density of 0.80 to 1.20 g/cm3 would be considered acceptable for most growing media.
Values considerably lower than these are commonly encountered with, artificial, soilless
growing media, and are just fine when it comes to plant growth.
Bulk density values of 1.25 - 1.65 g/cm3 have been shown to reduce root growth.
29
HORT 1001
Plant Propagation
Experimental Design
During the semester, you will be carrying out experiments, and reporting on them in written
form. Each experiment begins with experimental variables. What is an experimental variable and
how are they chosen? An experimental variable is defined as any individual characteristic that
has been chosen to be different from one group of individuals to another group within the
experiment. What does that mean for you in plant propagation?
For example, when you are attempting to root herbaceous stem cuttings, some examples of
variables that will be used during the semester are:




Effect of plant growth regulator application on inducing rooting - one group of cuttings
will be treated with one concentration of IBA and another group would not be treated.
Effect of treatment with different concentrations of a plant growth regulator - each group
of cuttings will be treated with different concentrations e.g. IBA at 1000, 3000, and 5000
ppm.
Effect of the carrier on plant growth regulator action - a group of cuttings treated with
IBA in talc vs IBA dissolved in alcohol.
Effect of leaf area on rooting success of cuttings - compare the effect of number of leaves
on rooting.
This list of possible variables could go on, limited only by time, space and supplies. However,
the variables could change depending on the type of plant material you were using. In
experimental design (at least for plant propagation class), there are 2 important rules to
remember when thinking about experiments: always use a control (plant material that remains
untreated); and always repeat your treatments.
What is a control?
A control is a group of cuttings that remain untreated. For example, if you were interested in
treating 5 cuttings of Spirea japonica with 1000 ppm IBA in talc, you would also leave 5
cuttings untreated. This allows you, at the end of the experiment, to conclude whether IBA at
1000 ppm in talc increased root initiation or increased the speed roots appeared. If you treated all
of the cuttings with 1000 ppm IBA you would have nothing to compare them to. The same goes
for all of the variables listed above. If you are going to treat a group of cuttings in a certain way,
you must leave the same number of cuttings untreated for the control.
What is a treatment?
A treatment is defined as the combination of variables applied to experimental units (each
individual cutting is an experimental unit) in the study. For single variable experiments, the
variable and the treatment are the same. For example, if you were comparing whether 1000 ppm
IBA enhanced rooting of Echniacea purpurea, the variable and treatment are the same – IBA
30
treatment. In an experiment comparing seed scarification treatments, the variable might be
scarification methods including the treatments mechanical, acid, and hot water scarification. All
other factors need to remain the same. So after treatment with different scarification methods, all
seeds would be planted at the same time, in the same media, placed side by side in the same
greenhouse. The only difference in the seeds would be the treatment that you imposed earlier.
You must take care when treatments involve combinations of several variables since the effects
can be confounded and make it impossible to make conclusions from the experiment. For this
reason, we will be doing very few multiple variable experiments.
Designing experiments
For each experiment designed, you begin by choosing your variables. How were the variable
chosen for the experiments that we are doing in Plant Propagation? We chose them to
demonstrate different methods to effect rooting in asexual propagation. We are experimenting
with plant material that we don’t know how successful some of the treatments will be. We
looked through the text and web sources to determine plant material that might work for the
treatments that we are imposing. The next step was to determine the number of experimental
units that will receive each treatment. For our example of herbaceous cuttings, each cutting
would be an experimental unit or for the scarification example each seed would be an
experimental unit. In this class, 5 experimental units should be sufficient but each treatment
should be repeated on no fewer than 3 experimental units. In a few experiments, each student
will only be doing one set of treatments and we will combine data from 5 or more students for
the final analysis.
Next, we thought about what kind of data should be collected. In the seed scarification
experiment you might collect data on number of seeds germinated and calculate the %
germination for each treatment; number of days to emergence and calculate the average for each
treatment. Other examples might be, number of roots, root length, days to rooting, number of
plantlets formed, etc.
Implementing your experiment
Collect and prepare the necessary plant materials and apply your treatments. Observe your
experiments frequently; at least once a week. Collect data as necessary. Carefully record exactly
what you did from beginning to end. Make sure you record dates in your lab book that you
started and ended the experiment as well as the date that you collected data.
Be sure everything is labeled! Labels must include the scientific name of the plant. Each
treatment needs to have its own label. You can use codes to identify treatments but be sure to
make notes confirming the meaning of the code. You will be doing lots of experiments and so
codes that were clear when you did them in September, might not be so clear after 12 more
experiments and 8 weeks!
When the experiment is completed and all data has been collected, transplant material you wish
to keep. Every pot that goes into the greenhouse must have a label or it will be thrown away. We
recommend taking these plants home regularly as we have limited greenhouse space and plants
don't do well being transported outside in December
31
Introduction to Experimentation:
Herbaceous Stem Cuttings
[E; individual; 1 species, 2 treatments; 5 replications/treatment = 10 cuttings/student]
Objectives:

To introduce experimentation including application of treatments (in this case treatment with
IBA) and data collection.
 To learn how to take cuttings.
 To produce uniformly rooted cuttings.
To Do Before Class:
Read p. 15 in your lab manual and pp. 154-156 in your lab text (Plant Propagation).
Plant Material:
Solenostemon scutellarioides [syn. Coleus  hybridus] (Coleus; Lamiaceae, Mint Family)
Treatments:
1. Control
2. 1000 ppm IBA in talc
Materials Provided:
 Pruners
 Razor blades
 Labels
 Permanent marking pen (black Sharpie)
Methods:
1.
2.
Label two colored plastic stakes with the date, treatment and the plant material (Coleus).
Take 10 cuttings of the plants provided, with each cutting having 2-3 nodes. Take each cutting just
below a node. The cuttings should all come from the same plant and be as similar as possible,
including size, and the location on the plant from which they were collected. Be sure to keep track of
polarity – which end was up – for each of your cuttings.
3. Wrap the cuttings in moist paper toweling.
4. Finish preparing your cuttings by removing the leaves from the bottom node of each cutting.
5. Once your cuttings have been prepared dip the bases of five (5) cuttings in 1000 ppm IBA in talc; the
remaining five (5) cuttings will be the controls (non-treated).
32
6. After treatment, take all 10 cuttings (keeping the treatments separate) and stick them into your mist
bench plot with the appropriate labels.
7. Check on your cuttings each lab period and record your observations. Begin by shaking off the sand
and wrap the cuttings in moist paper toweling, keeping treatments separate. Count the number of
cuttings that have produced roots. Then evaluate the cuttings on a 0-5 visual rating scale: 0 = no roots, 5
= roots abundant. In general, rooted cuttings are ready for harvest when they have a root system that is
about 33% to 50% the size of the top. The sample tables provided may be used to record your data.
9. When you are finished collecting your data, pot up all your rooted coleus cuttings; place them in the
designated location in the greenhouse for later use.
10. As always, be sure to clean up all work areas when you are finished. Good sanitation is extremely
important in greenhouse production systems so please be sure to do your part in keeping work and
growing areas clean throughout the semester.
Results – Data and Observations:
 Final data will be collected after 3 weeks. For this and subsequent reports your data should be
summarized for each date of observation and presented in the form of a bar graph or table.
Discussion – Using your results and observations, discuss your findings and answer
the following questions:

Where did the roots emerge from - between the node, at the node, or at the bottom of the cutting?
What is the significance of where the roots emerge?

Did the application of 1000 ppm IBA aid in rooting?

If you could change one thing about this experiment, what would it be and why?
Recommendation:

Based on your findings, provide a one-sentence recommendation outlining the best method of
propagating coleus using stem cuttings.
33
Example of Data Table – use one such table for each date data is taken
Title of Experiment: Introduction to Experimentation: Herbaceous Stem Cuttings
Date data taken: _______________
For the control:
# of cuttings started __________________
# of cuttings rooted __________________
Cuttings treated with 1000ppm IBA
# of cuttings started __________________
# of cuttings rooted __________________
Root Rating
Cutting Number
Treatment
1
2
3
4
5
Average
Control
1000 ppm
IBA
Note: This table is also provided as an example of the format that might be used when collecting
and reporting data for future experiment
Root ratings are an example of one type of data that will be collected for this and other
experiments. Use the diagrams below (Thanks to Ohio State University) as a guide to rate the
rooting success of your treatments. Assign a 1 to 5 rating to each cutting and use the data to
calculate an average root rating for each treatment.
34
Sexual Propagation: Using Scarification to Overcome
Hard Seed Coat Dormancy
[E; Individual; 2 species, 5 treatments, 2 replications = 20 seeds per student]
Objectives:

To experiment with different methods to overcome hard seed coat dormancy
To Do Before Class:
Read pp. 9-14 in your lab manual and pp. 19-20 (up to “Conditions needed for Germination”) in your
text.
Plant Material:
Gleditsia triacanthos (Common Honey Locust; Fabaceae, Bean Family)
Gymnocladus dioica (Kentucky Coffee Tree; Fabaceae, Bean Family)
Materials to Bring To Class:


Permanent marking pens
Lab notebooks
Materials Provided:





Files
Hammers
Labels
Pots
Potting medium
Treatments:
1.
2.
3.
4.
5.
Control (non-treated; seed coat not broken)
Mechanical scarification - hammer
Mechanical scarification - file
Hot water treatment
Acid scarification
Methods:
1. Make labels for each of the five (5) treatments – the two (2) species tested for each treatment will be
planted together in the same pot (= 4 seeds/pot).
2. Partially fill five (5) of the pots provided with growing medium. Each treatment will be planted in a
separate pot.
3. Plant two (2) control seeds for both species in one pot and label. The seeds should be planted such
that they are 1 to 2 times as deep as their diameter.
35
4. For the sake of timing and safety, seeds that have been soaked in boiling water and acid scarified
will be provided. Plant two (2) seeds of each species from each of these treatments together in
separate containers by treatment and label.
5. For one of the mechanical scarification treatments, scarify the seed coats of two (2) seeds of each
species with a file, making sure you file in the center of the seeds so you do not damage the
embryos. Once the seed coat is broken, stop so you do not damage the cotyledons. Plant seeds
and label.
6. For the second mechanical scarification treatment, scarify the seed coats with a hammer by
wrapping the seed securely in paper (this prevents the seed from shooting away when it is hit with
the hammer) and tapping until the seed coat is cracked. Ideally, pound hard enough to crack the
seed coat, but not so hard that you damage the seed. Plant the seeds and label.
7. Transfer the pots to the designated location in the greenhouse.
8. Check for germination each lab period. When you believe all the seeds that are going to germinate
have done so, you may want to dig up those seeds that did not germinate in an attempt to find out
why they failed to do so.
Results – Data and Observations:

Observations that may be taken for each treatment include (but are not limited to): % germination,
number of days to first emergence, type of germination (hypogeal or epigeal).
Discussion – Using your results and observations, discuss your findings and answer
the following questions:

What differences, if any, did you observe between treatments?

What are the advantages and disadvantages of using one scarification technique over another?
Recommendation:

Based on your findings, provide a one-sentence recommendation outlining the best method of
propagating these two species from seed.
36
Asexual Propagation: Adventitious Shoot Initiation in Response
to Treatment with BA
[E; individual; 1 species, 3 treatments, 5 replications/treatment
= 15 two-segmented phyllocad cuttings/student]
Objectives:
 To better understand the techniques involved in the propagation of new plants from pieces of the
parent plant; in this case phyllocads (a flattened stem or branch that functions as a leaf).
 To investigate whether treatment with BA influences the formation of adventitious shoot meristems
on phyllocad cuttings.
 To produce entire plants from phyllocad cuttings.
To Do Before Class:
Read pp. 4-6 and 15 (Herbaceous Stem Cuttings section) in your lab manual.
Plant Material:
Schlumbergera bridgesii (Christmas Cactus; Cactaceae, Cactus Family)
Materials To Bring To Class:
 Permanent marking pens
 Lab notebooks
Materials Provided:






Plant material
Razor blades
Label stakes
Newspaper
Benzyladenine solution (150 ppm in EtOH)
Benzyladenine solution (1500 ppm in EtOH)
Treatments:
1. Control (no treatment)
2. 150 ppm BA in 50% EtOH
3. 1500 ppm BA in 50% EtOH
37
Methods:
1. Select a plant from which to collect your cuttings.
2. Using a clean, sharp razor blade, collect fifteen (15) phyllocad cuttings (two phyllocad segments per
cutting) of similar size, age, and quality. Be sure to keep track of the polarity of your cuttings.
3. Divide your cuttings into three (3) groups of five (5) and prepare labels for each group indicating
treatment and date. One group will serve as a control. Treat the next two groups of cuttings in the
with BA (150 ppm & 1500ppm) by dipping the whole cutting into the solution provided. Be sure
to re-cover the solutions when not in immediate use to limit evaporation and associated increases in
concentration.
4. Stick your cuttings in your assigned plot in the mist bench along with the appropriate labels.
5. Observe your cuttings on a regular basis and record your observations.
6. The plants produced are yours to keep; when you are done, pot them up and take them home if
you like. If you have good quality plants and wish to donate any of them for use by students in
future classes, we would be happy to accept them. Please label all plants to be donated with their
full scientific and common names and the date.
7. As always, be sure to clean up all work areas when you are finished.
Results – Data & Observations:
 Data and observations that may be taken for each treatment include (but are not limited to): root
initiation, shoot initiation, subsequent root and shoot development, number of phyllocads
produced, etc.
Discussion – Using your results, discuss your findings and answer the following
questions:
 Which comes first, roots or shoots?
 From where do the roots and shoots arise?
 Are there differences between propagation by phyllocads and standard stem cuttings
 What effects, if any, did your treatments have on root and shoot development?
Recommendation:
 Based on your findings, provide a one-sentence recommendation outlining the best treatment for
propagating from Christmas cactus using phyllocad cuttings.
38
Asexual Propagation: Propagation of Plants Using
Leaf Cuttings
[E; Group; 1 species, 6 treatments, 3 replications/treatment =18 cuttings/student]
Objectives:

To observe effects of benzyl adenine (BA) on shoot meristem formation.

To produce entire plants from leaf cuttings.
To Do Before Class:
Read pp. 19 in your lab manual and p. 157 in your text.
Plant Material:
Saintpaulia ionantha (African Violet; Gesneriaceae, Gesneria Family)
Materials To Bring To Class:
 Permanent marking pens
 Lab notebooks
Materials Provided:





Plant material
Razor blades
Label stakes
Paper toweling
1500 ppm benzyladenine (BA)
Treatments:
Leaf-petiole cuttings
1.
Control
2.
150 ppm (BA)
Leaf-blade cuttings
39
1.
Control
150 ppm (BA)
2.
Leaf-piece cuttings
1.
Control
2.
150 ppm (BA)
Methods:
1. Remove 15 leaves from an African violet plant; 6 with petioles; 6 without petioles, and three that
will be cut down the midrib vein. You will have a total of 15 leaves. All cuttings should come from
the same plant and be as close to the same age and size as possible. You may want to arrange to
exchange with other students when the experiment is completed so you have a variety of African
violets to take home.
2. Keep leaves wrapped in a moist paper towel while working. Be gentle.
3. Treat the bases of three cuttings with 1500 ppm BA and leave 3 cuttings untreated for each
treatment. For the leaf-piece cuttings treat the midrib of 3 cuttings with the BA and leave three
untreated.
4. Stick the cuttings in your assigned plot in the mist bench. The cuttings should be oriented with just
the bases of either their petioles or leaf-blades stuck into the rooting media. For the leaf-piece
cuttings, the midrib should be in contact with the sand.
5. Observe your cuttings on a regular basis and record your observations. Lift the cuttings gently and
by lifting from beneath to avoid breaking any roots.
6. As your cuttings become well rooted, remove the cuttings from the mist bench, collect data, and pot
them up. Be sure to keep your plants labeled and continue to follow shoot development after
potting.
7. As always, be sure to clean up all work areas when you are finished.
Results – Data & Observations:
 Data and observations that may be taken for each treatment include (but are not limited to):
callus formation, root initiation and development, shoot initiation and development, time to
harvest and transplantation, plant (roots and shoots) quality, etc.
Discussion – Using your results and observations discuss your findings and answer
the following questions:
 Did roots and/or shoots initiate? If so, where?
 Which came first, roots or shoots?
 Were there any differences in root and shoot development between the treatments or among
40
the different types of cuttings?
 Why might a plant propagator use leaf-type cuttings compared to other types of cuttings (e.g.,
stem cuttings)?
 What effects, if any, did your treatments have on root and shoot development?
 If you could change one thing about this experiment, what would it be and why?
Recommendation:
 Provide a one-sentence recommendation outlining the best treatment from your data for
propagating African violet using leaf cuttings.
41
Sexual Propagation: Effects of Seed Coat Removal, Treatment
with GA3, and Cold Stratification on the Germination of Apple
Seeds
[GE; groups of 4; 1 species, 8 treatments, 5 seeds/treatment = 40 seeds/group]
Objectives:

To investigate the effects of handling and various pre-plant seed treatments on the
germination of apple seeds.
To Do Before Class:
Read pp. 4-6, 7-14 in your lab manual and pp. 10 & 19-20 (up to “Conditions needed for
Germination”) in your text.
Plant Material:
Malus domestica (Apple; Rosaceae, Rose Family) – cultivar announced in class.
Materials to Bring to Class:





Razor blades
Permanent marking pens
Ziploc bags
Tweezers
Lab notebooks
Materials Provided:





Label stakes
Paper toweling
Pots
Growing medium
GA
Treatments:
1.
2.
3.
4.
5.
6.
Stratify, Seed coats intact, No GA3
Stratify, Seed coats intact, GA3
Stratify, Seed coats removed, No GA3
Stratify, Seed coats removed, GA3
Direct Sow, Seed coats intact, No GA3
Direct Sow, Seed coats intact, GA3
42
7. Direct Sow, Seed coats removed, No GA3
8. Direct Sow, Seed coats removed, GA3
Methods (see flow chart; Figure 1):
1. For this experiment you will be working in groups of four (4).
2. Prepare label stakes for each of the treatments to be done today – Treatments 1-4. Be sure to write
the date and the names of your group members on each stake.
3. Fill five (4) pots or, if plug trays are used, four (4) rows of plugs with slightly moistened growing
medium.
4. Extract 25 apple seeds keeping them moist AT ALL TIMES by wrapping them in a moist paper
towel. Each treatment will be replicated five (5) times (5 seeds per treatment) for a total of 20
seeds. We are having you extract 25 seeds because one of the procedures, removing the seed coat,
may be difficult to do. Having extras is important!
5. Remove seed coats from 10 seeds – KEEP THEM MOIST!
6. Sow 5 seeds with seed coats in a pot or row of plugs and 5 without seed coats in another pot or set
of plugs (Treatments 1 and 3). Seeds should be planted at a uniform depth and should be no more
than 1 cm deep. Place the appropriate label stake in the growing container.
7. Place 5 seeds with seed coats and 5 without seed coats in a Petri dish filled with 400 ppm GA3, and
allow the seeds to soak for 1 hour (Treatments 2 and 4). Then sow the 5 GA3-treated seeds with
seed coats and the 5 GA3-treated seeds without seed coats into separate pots or set of plugs in the
plug tray. Place the appropriate stake in each pot or row.
8. Water carefully and place all pots or the plug tray in a large plastic bag with your group number and
names of each group member as well as the date of this portion of the experiment written with a
Sharpie. Place the bags into cold stratification (cool, moist conditions).
CONTINUING ACTIVITIES: Approximately 6 weeks after the
seeds were placed into conditions of stratification remove
the seeds from stratification:
1. Remove seeds prepared in the first half of the experiment from stratification.
2. Prepare label stakes for each of the treatments to be done today (Treatments 5-8). Be sure
to write the date and your group members name on the stakes.
3. Extract 25 apple seeds keeping them moist AT ALL TIMES. Each treatment will be replicated 5
times (5 seeds per treatment) for a total of 20 seeds. Again, we are having you extract 25 seeds
because it may be difficult to remove the seed coats.
4. Remove seed coats from 10 seeds – KEEP THEM MOIST!
5. Sow 5 seeds with seed coats and 5 seeds without seed coats into separate pots or rows of the plug
43
tray (Treatments 5 and 7). As before, pay close attention to depth of planting. Label with the
appropriate stakes.
6. Place 5 seeds with seed coats and 5 seeds without seed coats in a Petri dish filled with 400 ppm
GA3, and allow the seeds to soak for 1 hour (Treatments 6 and 8). Then sow seeds into 2 separate
pots or rows of the plug tray. Label with appropriate stakes.
7. You will have 8 pots or 8 rows of a plug tray total (each with one treatment). Place the pots or plug
trays in the greenhouse.
8. Observations on germination must be taken and recorded each lab period. You may make your
observations as a group or you can designate one member to collect the data and report to the
group.
Results – Data and Observations:

Data and observations that may be taken for each treatment include (but are not limited to): # days
to emergence, # of seeds rotted, germination % (# seeds germinated/total number of seeds),
seedling size and vigor. You should discuss data collection as a group to standardize your efforts.
Also, at the end of these lab directions there is a data sheet that we will be using in the lecture
portion of this class. Information for the data sheet should come from lab observations.
Discussion – Using your results and observations discuss your findings and answer
the following questions:

How did the treatments compare with respect to germination percentage?

Is there a difference in germination if seeds are stratified vs. direct sow?

What effect does stratification have on germination?

Is removing the seed coat analogous to scarification?

Are there inhibitors to seed germination in the seed coat? How might you test your ideas?

What other treatments might be used to overcome dormancy requirements?

What recommendations would you give to someone who wanted to know about overcoming
dormancy of woody species?
Recommendation:

Based on your findings, provide a one-sentence recommendation outlining the best treatment for
germinating apple seeds.
44
Apple Seed Experiments - Data Sheet:
Treatment
# of Seeds Germinated
Stratified
1. With seed coat
2. With seed coat, GA3 treated
3. Without seed coat
4. Without seed coat, GA3 treated
Not Stratified
5. With Seed coat
6. With seed coat, GA3 treated
7. Without seed coat
8. Without seed coat, GA3 treated
Comments:
45
Date:
% Germination
Figure 1. Flowchart of Apple Seed Experiments
with GA
Treatments 2 & 4
without GA
Treatments 1 & 3
Stratification
(with & without seed coat)
with GA
Treatments 6 & 8
No Stratification
(with & without seed coat)
without GA
46
Treatments 5 & 7
Sexual Propagation: Effects of Growing Medium on Seed
Germination and Seedling Performance
[E; 1 species, 7 treatments 3 replications (3 containers of each medium; 4
seeds/container) = 21 containers and 84 seeds]
Objectives:
 To demonstrate the effects of various growing media on seed germination and seedling
performance.
 To highlight the importance of selecting a suitable growing medium based on the plant being grown
and the container used.
To Do Before Class:
Read pp. 28-29 in your lab manual and pp. 32-35 in your text. On web: Growing media in container
production in greenhouse and nursery. Part 1: Components and Mixes
http://www.uaex.edu/Other_Areas/publications/PDF/FSA-6097.pdf &
http://www.uaex.edu/Other_Areas/publications/PDF/FSA-6098.pdf
Plant Material:
Tagetes sp. (Marigold; Asteraceae, Aster/Sunflower Family)
Materials To Bring To Class:
 Permanent marking pens
 Lab notebooks
Materials Provided:




Containers
Various growing media (use the same media that were used for the porosity/bulk density experiment)
Marigold seeds
Label stakes
Treatments:
1.
2.
3.
4.
5.
6.
7.
Growing Medium #1
Growing Medium #2
Growing Medium #3
Growing Medium #4
Growing Medium #5
Growing Medium #6
Growing Medium #7
47
Methods:
1. This experiment will be done in groups; the same groups as for the porosity/bulk density
experiment.
2. Fill three (3) containers to 1/4" below the brim with each of the seven (7) growing media. for a total
of 21 containers filled with growing media.
3. Each of us will fill a container with medium differently. For example, I might pack the medium
more tightly and have a different understanding of what full means than someone else does. To
reduce the effects such differences in the method of filling containers might have on the
experimental results, the same person should fill all the containers with the appropriate media.
4. Label each container indicating your group name, the date, and the medium.
5. Plant 4 marigold seeds on the surface of the medium in each container.
6. Cover the seed with additional medium filling each container to the brim resulting in a planting
depth of 1/4". Be sure to cover the seed with the same medium as is already in the container and
once again, have the same person perform this task (this does not have to be the same person who
initially filled the containers).
7. Carefully water your seeds; wet the medium completely being very careful not to splash medium
from the containers or disturb your seeds.
8. Place your containers in the flats provided and carefully transfer them to the designated location in
the greenhouse.
9. Check your seeds and follow the germination and growth of your seedlings each lab period until
final data is collected. Water your containers carefully as needed.
10. As always, be sure to clean up all work areas when you are finished.
Results – Data & Observations:
 Data and observations that may be taken for each treatment include (but are not limited to): time to
emergence of first and last seedlings, % germination, seedling vigor, seedling appearance, and %
survival.
Discussion – Using your results, discuss your findings and answer the following
questions:
 Did your different media influence seed germination? If so, why?
 Was the growth of your seedlings influenced by the various media? If so, how?
 Which medium was the best choice for growing marigolds from seed to the seedling stage in the
containers used?
48
 How do your results match up with your earlier hypotheses about the suitability of each medium
for growing plants based on porosity values?
Recommendation:
 Based on your findings, provide a one sentence recommendation outlining the best medium for
growing marigolds in the containers used
49
Asexual Propagation: Rooting of Herbaceous Stem Cuttings
in Response to Type of Auxin, Concentration of Auxin,
Different Carriers of Auxin
[P; 1 species, each student will do one experiment, with 4-6 treatments per experiment,
5 replications/treatment = 20-30 cuttings/student; shared data for the final report]
Objectives:
 To successfully propagate plants from stem cuttings.
 To observe how treatment with auxin influences rooting in different plants.
To Do Before Class:
Read pp. 4-6 and 15 in your lab manual and pp. 154-155 in your text.
Plant Material:
The plant material used will be determined by availability in the horticulture garden.
Materials To Bring To Class:




Pruners
Permanent marking pens
Polyethylene bags
Lab notebooks
Materials Provided:
 Label stakes
 Paper toweling
 Treatment compounds
50
Treatments:
Experiment 1 - Rooting in Response to Different Types of Auxin
1.
2.
3.
4.
5.
Control
50% ethanol
1000 ppm IBA in 50% ethanol
1000 ppm NAA in 50% ethanol
1000 ppm IAA in 50% ethanol
Experiment 2 - Rooting in Response to Different Concentrations of an Auxin
1.
2.
3.
4.
5.
Control
1000 ppm (0.1%) IBA in talc
3000 ppm (0.3%) IBA in talc
8000 ppm (0.8%) IBA in talc
16,000 ppm (1.6%) IBA in talc
Experiment 3 - Rooting in Response to Different Carriers of an Auxin
1.
2.
3.
4.
Control
50% ethanol
1000 ppm IBA in 50% ethanol
Talc
5. 1000 ppm IBA in talc
6. 1000 ppm IBA (K-IBA) in water
Methods:
1. Experiments will be assigned by the instructor. Each student will do one experiment, selecting one
species for their cuttings. Be sure to record the genus, species, and common name of the plant.
2. Each student should prepare labels for each of the treatments included in their experiment. The
stakes should also be labeled with the date, the name of the plant, and a brief description of the
experiment (i.e., type, concentration, carrier).
3. Each student should collect the required number of cuttings needed for their experiment from the
plant species chosen. In all cases, the cuttings should have four (4) nodes with fully expanded
leaves (ask if you are unsure how to do this). As you collect your cuttings, be sure to place them in
polyethylene bags with moist paper toweling to prevent them from drying out.
4. Return to the classroom and finish preparing the cuttings by removing the leaves from the bases of
the cuttings. Keep the cuttings covered with moist paper towel to keep them from drying out.
Divide the cuttings into groups of five with one group of five cuttings for each of the treatments in
your experiment. Treat the cuttings in each group with one of the treatments included in your
experiment.
5. After treatment, carefully transfer the cuttings to the mist house and stick them in your plot in the
51
mist bench with the corresponding label. Keep your cuttings separated and labeled by treatment at
all times.
6. As always, be sure to clean up all work areas when you are finished.
7.Observe the cuttings for each of the experiments at least weekly and record your observations.
Results – Data & Observations:
 Data and observations that may be taken for each treatment in each experiment include (but are
not limited to): callus formation, rooting (time to first root emergence, time to harvest, % rooting,
and root system quality and symmetry), signs of necrosis at the site of application, length of longest
root, # of roots, and performance of the cutting growth.
When taking data, pay close attention to where roots are forming.
Discussion – Using your results and observations, discuss your findings and answer
the following questions:
 Where along the stem were roots initiated?
 Did rooting continue to improve over time?
 What effects, if any, did your treatments have on rooting?
 Which treatment(s) resulted in the best rooting?
 If you could change one thing about this experiment, what would it be and why?
Recommendation:

Discuss your observations with other members of your group and, as a group, provide a onesentence recommendation for each experiment outlining the best treatment for propagating your
specific herbaceous plant using stem cuttings.
52
Asexual Propagation: Designing an Experiment to Maximize
Rooting of Leaf-Petiole and Leaf-Bud Cuttings
[P; individual; 1 species; experiments designed individually by each student using
a minimum of 20 experimental units]
Objectives:

To review the scientific method by providing an opportunity to design an experiment involving leafbud and leaf-petiole cuttings.

To compare the effects of various treatments on leaf-bud and leaf-petiole cuttings.
To Do Before Class:
Design your experiment by clearly identifying objectives, treatments to be done, and data to be taken.
You need to use at least 20 experimental units. You must choose the variables. Each of the resulting
treatment combinations should be replicated five (5) times. You must have your experimental design
approved by the instructor prior to initiating your experiment.
Read pp. 4-6, pp. 19 and 30-31 in your lab manual.
Plant Material:
Peperomia scandens ‘Variegata’ (Variegated False Philodendron; Piperaceae, Pepper (the spice)
Family)
Materials To Bring To Class:



Pruners
Permanent marking pens
Lab notebooks
Materials Provided:








Plant material
Label stakes
IBA in talc (1000, 3000, 8000, 16000 ppm)
IBA in water (1000 ppm)
IBA in 50% ethanol (1000 ppm)
NAA in 50% ethanol (1000 ppm)
BA in 50% ethanol (150ppm & 1500 ppm )
Distilled water
53
 talc
 50% ethaol (ETOH)
Treatments:
To be determined by student – leaf-bud and Leaf-Petiole cuttings combined with other experimental
variables selected by the student.
Methods:
To be determined by student.
Results – Data & Observations:
 To be determined by the student - data and observations that may be taken for each treatment
might include (but are not limited to): root initiation (time to rooting and root number), root
quality and percentage, shoot initiation (time to shoot initiation and shoot number), subsequent
aspects of root and shoot development, etc.
Discussion – Using your results, discuss your findings and answer the following
questions:
 Which came first roots or shoots?
 Where did the roots and shoots arise?
 Were multiple or single plants produced?
 Were there differences between the treatments when compared?
 Were there differences between types of cuttings?
Recommendation:
 Based on your findings, provide a one-sentence recommendation outlining the best method for
propagating Peperomia scandens ‘Variegata’ from leaf-bud cuttings and a one-sentence
recommendation outlining the best method for propagating Peperomia scandens ‘Variegata’
from leaf-petiole cuttings?
54
Sexual Propagation: Manipulation and Germination of Seeds
from an Herbaceous Perennial, or Woody
Perennial Plant
[P; individual; 1 species of choice, treatments and replication numbers to
be determined by the student]
Objectives:
 To determine the requirements for successfully germinating seeds from a variety of plants based on
research and experimentation.
 To experiment with different techniques used to break seed dormancy.
 To design your own experiments to answer specific, testable research questions pertaining to seed
handling and germination.
To Do Before Class:
Read the Introductory Information section on seeds in the lab manual (pp. 7-14).
You will need to do some research to hopefully learn about the germination requirements of the plant you
are working with before you set up the experiments. Don’t put this off! A little research will go a long way
when it come to success or failure with this project. There are many books and research papers specific to
the germination of seeds from various plants. The internet can also be a valuable source of information.
One website that has information on many garden plants is the Plantfacts site at http://plantfacts.osu.edu/.
And then, of course, you can always visit the library.
Plant Material:
Seeds of one type of plant material (1 herbaceous perennial or 1 woody perennial) to be distributed by the
lab instructor.
Materials To Bring To Class:
 Permanent marking pens
 Lab notebooks
55
Materials Provided:







Label stakes
Paper toweling
TTC (for viability testing)
Gibberellic acid (400 ppm)
Containers
Growing media
Scarification and stratification facilities
Treatments:
To be determined by each student depending on the plant material chosen.
Methods:
To be determined by each student depending on the plant material chosen. This may require several
attempts and experimentation to achieve successful results.
Results – Data and Observations:

Observations that may be taken for each treatment may include (but are not limited to): number of
days to germination, % germination, % viable seed, type of germination (epigeal or hypogeal),
monocot or dicot, type of dormancy, etc.
Discussion – Using your results and observations discuss your findings in the
following format and answer the following questions:

!
Don’t worry if you are not successful in germinating the seeds you have been given; the point is
that you make a reasonable attempt. If you have a problem getting seeds to germinate, you should
consider and develop a hypothesis as to why you think you were unsuccessful. Again, research into
the germination requirements of your seeds is necessary in this regard.
Based on the information gained while attempting to germinate your seeds and research on your
selected plants, you should design a seed packet for each species of plant that you collected and
researched. As an example, think of the vegetable and flower seed packets you have seen offered for
sale. Be creative in your seed packet designs; remember marketing is an important aspect of
56
any successful horticultural enterprise. What type of design and information would catch your
attention and persuade you to purchase a packet of seeds?
At a minimum, each seed packet should include the following information:
1.)
A picture or drawing of the mature plant and a young seedling.
2.)
Family relationship (family name), scientific/botanical name (Genus species, and variety if
applicable), and common name(s).
3.)
Classification based on lifecycle (herbaceous perennial, woody perennial tree or shrub) and
descriptive information (mature height, habit, etc.).
4.)
Geographic range of the species (Where is it native?).
5.)
Cultural information – planting directions, light requirements (sun/shade), moisture
requirements, soil preference (texture, pH), fertility requirements, preferred habitat,
potential pests, etc.
6.)
Specific information gleaned from your research and experiments – number of days from
sowing to emergence, % viable seed, % germination, germination requirements (light,
temperature requirements), treatments needed to overcome dormancy, etc.
7.)
Price – How much are your seeds worth; how many seeds/packet and how much would
you charge?
8.)
References used (at least 3 references should be used – books, periodicals, and the internet
–the internet cannot be the only source for references).
Recommendation:

Based on your findings, provide a one-sentence recommendation outlining the best method for
germinating the seeds for each of the species you investigated.
57
Lets Go Grocery Shopping: Propagating Produce
[P; Individual; at least 3 species; 5 propagation methods; at least 2 experimental treatments/
technique]
Although you may have never thought of it as such, your neighborhood grocery store may serve as a rich
source of propagules for the production of new plants at home. For this project, your objective will be to
successfully propagate new plants from produce purchased in local grocery stores. Proceed by selecting the
produce you intend to work with and then formulating a strategy to propagate new plants from your
purchases. Experiment with methods that are different than the standard practice; in fact for many types of
produce, based on what is available in stores, you will have no choice but to use a different method. For
example, cole crops such as cabbage or brussel sprouts are commercially propagated from seed, however,
you would need to determine how to propagate them from the heads of cabbage or the individual sprouts
or stalks sold in stores. Design experiments to determine the best method of propagating your material
(e.g., different types of propagules or stem cuttings with and without auxin treatment) and/or trying similar
techniques on a variety of crops. Be creative and don't be afraid to take some chances just for the fun of it.
Your goal is to produce multiple plants from one source. When considering and choosing propagation
techniques, remember this goal!
Some propagation techniques and potential treatments to consider include:
 Seeds (but not prepackaged vegetable or flower seeds; sorry! Seeds count as one technique);
treatments might include various treatment to enhance germination.
 Scooping, Scoring, Chipping, or Scaling of bulbs to produce bulblets; treatments might include
method (e.g., scooping vs. scoring) or treatment with BA (Note – this can be a tricky one; be sure what you
think is a bulb, is really a bulb.).
 Leaf cuttings (leaf-piece, leaf-blade, leaf-petiole, or leaf-bud cuttings); treatments might include type
of cutting or growth regulator treatments.
 Stem Cuttings (herbaceous stem or softwood, semi-hardwood, or hardwood cuttings); treatments
might include size of cutting, wounding, or treatment with growth regulators.
 Root Cuttings; treatments might include size of cutting or treatment with growth regulators (Note – this
can also be a tricky one; be sure what you think is a root really is root tissue. ).
 Division (tubers, tuberous roots, tuberous stems, rhizomes); treatments might include different
growing media or size of cutting.
 Grafting
58
 Other Techniques; check with the instructor if you have other ideas.
To satisfy the requirements of this laboratory exercise, you are required to:
1. Experiment with at least five (5) different propagation techniques.
2. Apply at least two experimental treatments per technique; one treatment should be a control. If
you attempt grafting as a technique, you will not be required to apply treatments in that case.
3. Successfully propagate completely new plants (roots and shoots) with at least one (1) technique.
4. Work with at least three (3) different species of produce. The five techniques you choose may
be applied to the same or different varieties of produce (e.g., both leaf and stem cuttings from the
same species or leaf cuttings from one species and stem cuttings from another). Remember,
seeds count as one technique so propagating three species from seed doesn’t count as three
techniques. Once again, be creative in your plant selections and methodology. Anyone can
easily propagate potatoes from the tubers available in stores, and potatoes are certainly a fine
choice as one selection; however, you are encouraged to try more unusual produce selections in
your trials.
When making your produce selections, be sure to think about what techniques might apply. Not
all choices will work; bananas are a good example – Why?
5. Cite at least three references, one of which must be from a source that is not web based.
In addition to achieving at least a modicum of success, to receive full credit, your methods must be clearly
deliberative and appropriate to the produce you have selected. For example, dried bay leaves are obviously
dead, and no matter what the methods used, an unsuccessful outcome is predetermined. Also, growth
regulator treatments should be appropriate for the technique used.
Remember, propagation entails the efficient and economical production of many plants using propagules
collected from a group of existing plants or perhaps even from a single mother plant – in other words
propagation involves multiplication. It is not simply growing a single onion plant from an onion bulb, which
is neither efficient nor economical. Multiplication - optimal increases in the number of quality plants in
production - is key to any successful propagation strategy.
Your efforts to propagate your produce selections will be presented in poster form. Posters are a common
and concise method used by researchers to present research findings at scientific meetings. Feel free to be
creative as you design your poster. It should be complete, yet brief and to the point. The poster should
have a title. Be sure the species, techniques, and treatments used are clearly identified; you should
essentially present a mini lab report for each of the experiments performed. As a general outline, you might
consider using your produce selections as the primary sections followed by the experiments performed
including the techniques and treatments used. For each produce selection, the following information
should be presented:
1. Objective(s).
2. Plant material used – identify the produce selections using their correct botanical (Genus, species,
59
and cultivar if known) and common names. You should also indicate what plant families your
produce selections belong to.
3. Methods used – describe the techniques used and the treatments applied.
4. Results – success or failure; if you were unsuccessful, indicate why you think your efforts failed.
5. A description of how your produce selections are typically propagated commercially.
6. A one sentence recommendation – based on your efforts, what was the best way found to
propagate your produce selections; if you were not successful, indicate what you might
recommend trying next.
7. Literature cited – list the references from which you obtained information about your produce
selections.
Posters will be presented during a show-and-tell session at the end of the semester. Samples of your
successes should be part of your presentation.
60
Sexual Propagation: Seed Morphology, Viability Testing,
Germination Testing, and Epigeal/Epigeous vs.
Hypogeal/Hypogeous Germination
[T; individual; Seed morphology 4 species, 2 replications per species = 8 imbibed seeds/student,
Seed viability 4 species, 5 replications per species = 20 imbibed seeds/student
Hypogeal vs. Epigeal, 4 species, 5 replications per species = 20 dry seeds/student]
Objectives:
 To test viability of different seeds.
 To use different methods for determining germination percentage.
 To observe different types of germination.
To Do Before Class:
Read pp. 7-14 in your lab manual and p. 20 in your text - “How a Seed Germinates”.
Review the information on cut tests and tetrazolium staining at the following web site:
http://msucares.com/pubs/publications/p1978.htm
Plant Material:
Allium cepa (Onion; Liliaceae, Lily Family)
Phaseolus vulgaris (Bean; Fabaceae, Bean Family)
Pisum sativum (Pea; Fabaceae, Bean Family)
Zea mays (Corn; Poaceae, Grass Family)
Materials To Bring To Class:
 Permanent marking pens
 Polyethylene bags
 Lab notebooks
Materials Provided:
 Labels
61





TTC
Growing medium
Pots
Presoaked seeds
Dry seeds
Methods:
I. Seed Morphology
a. Obtain two (2) presoaked (imbibed) seeds and one (1) dry seed for each of the four species
provided. The presoaked seed have been soaked in water overnight to allow them to absorb
water. The uptake of water by living seeds is called imbibition and the seeds are said to be
imbibed. Imbibition is the first step in seed germination. First, observe the differences between
the seeds of the different species. Then compare the dry seed to the imbibed seeds for each
species. Diagram and label your seeds and make notes about your observations. When you
have finished observing your seeds from the outside, carefully disect them open and diagram
your observations. Microscopes will be available.
II. Viability Testing (% viable seed)
a. Cut tests allow you to look inside the seed to observe the health of the embryo.
i. Select five (5) presoaked seeds for each of the four species for a total of 20 seeds.
ii. Work on moist a paper towel, cut the corn and onion seeds in half with a sharp razor blade.
Remove the seed coats from the beans and peas and pull the halves apart.
iii. Observe the condition of the seed and the color of the embryo for each seed. As a general
rule, if the embryo is white in color, it is probably viable.
iv. Record your observations.
b. Tetrazolium tests (TTC) utilize an enzyme reaction in which live tissues stain red, and dead
tissues do not stain. TTC tests can be used to estimate seed germination and vigor and can be a
useful tool in determining seed quality and viability during harvesting, conditioning, storage, and
distribution.
i. Place the seeds that you cut open in a petri dish, cover with tetrazolium (TTC) solution, and
let sit for up to 1 hour checking staining intensity every 15 minutes.
ii. After you have reached the desired level of staining, drain the tetrazolium solution and rinse
the seeds two to three times in cold tap water and evaluate immediately.
iii. Evaluate the stained seeds under magnification and good light. The most desirable color for
seeds is a dark pink to a light red. Darker red seeds may also be viable. Viable seeds may be
completely stained, or the seeds may have slight, small dead areas over the cotyledon or the
chalazal end, or have dead or weak tissue over less than one-third of the cotyledonary area.
The radicle tip in a good seed may be quite dark, because this is an area of high-metabolic
62
activity, and the small amount of tissue allows deeper penetration of the tetrazolium solution,
resulting in a dark appearance.
III. Determining type of germination - epigeal/epigeous vs. hypogeal/hypogeous germination
Epigeal germination = hypocotyl elongates and the cotyledons rise above the soil
surface
Hypogeal germination = epicotyl elongates and the cotyledons remain below the soil
surface
a) Plant five (5) seeds of each species in a pot filled with growing medium. You will have 4 pots,
one for each species. Label the pot with a stake (marked with the species, your name, and the
date) and place in the greenhouse. Also wrap (5) seeds of each species in a damp but not wet
paper towel. Place the paper towel with the seeds in a plastic bag and store at room temperature.
b) Observe these seeds each lab period. As they germinate, compare your observations to those
for seeds germinated in rag dolls to confirm your conclusions. When it comes to determining
whether germination is epigeal or hypogeal, it is sometimes difficult to visualize what is
happening from seedlings germinated in rag dolls. It can be difficult to discern what the
orientation and relationship of the seedling to the soil would be had it actually been planted.
c) After your seeds in the rag dolls have germinated, carefully examine and diagram your seedlings.
Compare your observations to those made earlier. Based on your observations, determine
germination percentage, whether germination is hypogeal or epigeal, and whether the plant is a
monocot or a dicot, for each type of seed.
d) Follow the emergence of the seeds planted in pots and compare to the seeds germinated in rag
dolls to better understand the difference between epigeal and hypogeal germination.
Observations:
 Observations for each species, should include (but not be limited to): % germination, rate of
germination, type of germination (hypogeal or epigeal), and whether the species is a monocot or a
dicot.
Some Questions for Thought:
 What environmental factors may have influenced the rate of germination?
 Which technique for determining viability is the easiest? Fastest? Gives the most information?
 If you were setting up a seed testing laboratory, which of these techniques might it be important to
include? Why?
63
Asexual Propagation: Whip & Tongue Grafting
[T; individual; 5 grafts/student]
Objectives:
 To better understand the theory behind and demonstrate the techniques involved in whip & tongue
grafting.
 To successfully propagate plants using whip & tongue grafts or at least come to appreciate the skills
involved in grafting as a method of propagation.
 To successfully complete four whip & tongue grafts without cutting oneself; please be careful!
To Do Before Class:
Read pp. 23-25 (up to “Budding”) in your lab manual and the sections on grafting in general
and whip & tongue grafting in your text (pp. 58-59).
Plant Materials:
Cornus sericea (Red-twig or Red-Osier Dogwood; Cornaceae, Dogwood Family) = stock or scion
Cornus sericea 'Flaviramea' (Yellow-twig Dogwood; Cornaceae, Dogwood Family) = stock or scion
Materials To Bring To Class:
 Pruners
 Lab notebooks
Materials Provided:






Budding rubbers
Parafilm
Label stakes
Paper toweling
Peat moss (moistened)
Rooting promoter (3000 ppm IBA talc)
64
Methods:
 We will be using red-twig and yellow-twig dogwood stems to create our whip & tongue grafts. By
grafting these two dogwood selections with different colored stems together, we will be attempting to
create a novel or specialty plant that has both red and yellow winter stems. Grafting together two
plants that look very different will also serve to visualize and illustrate the grafting process.
1. Prepare the stock:
a. We will be using unrooted, hardwood cuttings for our rootstocks. The scion piece will be
grafted onto the unrooted cutting. The cutting will then be rooted at the same time the graft
heals. Once again this technique is called stenting.
b. Select enough woody stems of either red-twig or yellow-twig dogwood to prepare four (4)
cuttings. Stems that are pencil-sized or slightly larger are best. You can use all red-twig or all
yellow-twig for your rootstocks or you might want to try using a couple of each as stock pieces.
Each cutting should have at least one (1) node at its base and a relatively long section of
internode above. Depending on the length of the internodes for the plant material available,
your rootstock cuttings may have more than one (1) node.
Note: You may want to collect your scion wood at the same time to ensure that you have stems
of equal diameter (see 2. a. below).
c. Make the grafting cuts (see Figure 1). The cuts required for the whip & tongue graft are fairly
simple and straightforward. Using a clean sharp razor blade, begin by making a slanting cut 1 to
1.5 inches (4cm) long within the internode section at the top of each rootstock cutting. Start at
a point about ½ the distance between the pith and the outer edge of the bark, on the upper side
of the slanting cut, make a second cut straight downward about half as long as the first cut to
form the “tongue”.
d. To insure a good fit and close contact between the stock and the scion pieces, all cuts should be
straight and clean; remember, too, that hemoglobin is not effective as a plant growth regulator.
Don’t let your wood dry out; keep your stock pieces wrapped in moist paper toweling in a
polyethylene bag when you are not working with them directly and be aware that the wood will
dry out as you are making your graft cuts.
2. Prepare the scion:
a. Select just enough woody stems to make four (4) scions having opposite stem colors as the
stock pieces already prepared. It is important that the wood used for the scions be equal in
diameter to the stock pieces. Each scion piece should have one (1) or two (2) pairs of buds
(nodes) at the top and a relatively long section of internode at its base.
b. Make the grafting cuts (see Figure 1). Holding the scion piece upside down, make exactly the
same cuts in the base of the scion piece as were made in the stock; the length and angle of the
cuts on the scion piece must be identical to those on the stock to insure a good match. Again,
keep your scion pieces from drying out.
3. Join the stock and scion pieces:
65
a. Slide the stock and scion together. It is very important that the stock and scion pieces fit
together tightly (See Figure 1). Hold the grafts up to the light to look for gaps and make
adjustments as needed. Once again, keep your stock and scion pieces, especially the cut
surfaces that will be joined together, from drying out.
b. Wrap the joined pieces with parafilm followed by a budding rubber. Wrap the budding rubber
from the bottom up. Both the parafilm and the budding rubber should extend a half inch or so
above and below the cut surfaces of the graft to provide sufficient support. A budding rubber
may be used alone, however, the parafilm is easier to apply and helps reduce frustration by
holding the pieces of the graft in place while the budding rubber is applied. Do not use
parafilm alone as it breaks down and does not last long enough.
c. Don't get frustrated. Remember, learning to graft takes practice and patience. Your technique
should improve with each attempt.
4. Treat the bases of your grafted cuttings with 3000 ppm IBA in talc.
5. Stick two (2) of your completed grafts directly into your plot in the mist bench. Be sure to label
them. Pre-callus the remaining two (2) grafts for two weeks prior to sticking by packing them in
moist peat moss in a polyethylene bag; label the bag with your name and the date and place in the
box provided.
6. Observe your grafts on a regular basis and record your observations. Avoid the temptation to
remove the budding rubber and parafilm to "sneak a peek" for at least five (5) weeks.
7. When your grafted rootstocks are well rooted, and the grafts have healed, pot them up. The grafts
will continue to heal in the greenhouse. If your grafts should show signs of wilting after potting,
enclosing the potted graft in a polyethylene bag for a time and then gradually hardening-off may
be helpful.
8. As always, be sure to clean up all work areas when you are finished.
Observations:
 Observations should include (but not be limited to): callus formation, bud break, root development
and success or failure of your grafts.
Some Questions To Ponder:
 Were your grafts successful or not? If not, why not?
 What factors might have influenced the success or failure of your whip & tongue grafts?
 Based on this experience, what might you do differently in subsequent attempts at whip & tongue
grafting?
 Why is it important for the stock and scion to be the same diameter? What if they were not
similar?
66
 Why is grafting done? Why do you suppose grafting is a more expensive means of propagation?
 Which factor would you say is most important in grafting success?
67
Figure 1. Making a Whip and Tongue Graft.
Preparing
Preparing
the scion
Joining the stock and
the stock
the scion
68
Asexual Propagation: Unique Vegetative Structures
[T; individual]
Objectives:

To become familiar with storage structures

To better understand and demonstrate several techniques commonly used to propagate plants with
unique vegetative storage structures.
To Do Before Class:






Read 21-22 in your lab manual, 25-26 (pictures on p. 27) in your text
Scaling p 258 in your text
Rhizomes pp. 149, 169, 184, 191, and 288 in text
Stolons p162-163 in text
Bulbils p.273 in text
Tubers p. 235 in text
Types of structures and plant material to be used:
Scaly Bulbs: Lilium sp. (Asiatic Lily; Liliaceae, Lily Family)
Tunicate Bulbs: Tulipa Sp. (Tulip; Liliaceae, Lily Family)
Rhizomes: to be determined
Stolons: spider plant, strawberries
Bulbils: Tiger lily (Lilium lancifolium, Liliaceae, Lily Family)
Tubers: Potato
Materials To Bring To Class:



Pruners
Latex gloves (if you think you might be sensitive to plant exudates)
Lab notebooks
Materials Provided:
 Pots
 Labels
 Incubation medium – 50:50 milled sphagnum moss:perlite
69
Methods:
For the purpose of comparison, and to better understand their structure, observe and diagram the dissected
tunicate (tulip and grape hyacinth) and scaly (lily) bulbs provided. Other types of “bulbs” (corms, tubers,
tuberous roots, etc.) may also be provided for the purpose of observation. Be sure to record your
observations and label your diagrams.
Scaly Bulbs
1. Obtain structures for propagating:
 10 lily scales
 3 pieces of rhizome
 3 pieces of stolon
 2 bulbils
 1 tuber
2. The shared lily bulb will be used to demonstrate scaling (see p. 258 in your text). Simply separate
10 individual scales from the basal plate of your lily bulb. The scales should be plump and in good
condition.
3. Place your bulb scales into a clean polyethylene bag together with three to four times their volume
of the moistened incubation medium provided.
4. Be sure to label the bag with your name, the date, the species, and the method used. Place your
bag in the boxes provided; they will be incubated in a warm, dark place.
5. Your bulb scales will be available for observation during subsequent lab periods during the
semester; observe them on a regular basis for bulblet development and record your observations.
Remove any materials showing signs of decay; if you are not sure, ask.
Rhizomes: To be determined
1. Remove plants from pots if necessary. Using a sharp razor blade cut off a piece of the rhizome (see
pictures p. 288 in your text).
2. Each cutting needs to have at least one node on it.
3. Plant the pieces of rhizome together in a pot, cover with soil. Label your pot with name, date, type
of structure and place it in the greenhouse.
4. In subsequent lab periods, take observations on above ground growth.
Stolons: spider plant, strawberries
1. Remove plants from pots if necessary. Using a sharp razor blade, cut off a piece of the stolon (see
pictures p. 197 in your text, they are called runners).
2. Each cutting needs one node.
3. Plant the pieces of stolon together in a pot, cover with soil. Label your pot with name, date, type of
70
structure and place it in the greenhouse.
4. In subsequent lab periods, take observations on above ground growth.
Bulbils: Tiger lily (Lilium lancifolium, Liliaceae, Lily Family)
1. Remove 2 bulbils from plants (see pictures p. 273 in your text).
2. Plant the 2 bulbils in a pot, cover with soil. Label your pot with name, date, type of structure and
place it in the greenhouse.
3. In subsequent lab periods, take observations on above ground growth.
Tubers
1. Remove plants from pots if necessary. Using a sharp razor blade, divide tuber (see pictures p. 235
in your text).
2. Each tuber cutting needs to have at least one node on it.
3. Plant the pieces of tuber together in a pot, cover with soil. Label your pot with name, date, type of
structure and place it in the greenhouse.
4. In subsequent lab periods, take observations on above ground growth.
Wash your hands and, as always, be sure to clean up all work areas when you are finished.
Observations:
 Observations should include (but not be limited to): labeled diagrams of the various bulbs
provided, the success or failure of each technique, time to bulblet initiation, and the number and
relative size of the bulblets produced.
Some Questions to Ponder:
 For each species, which method of propagation, scooping, scoring, chipping or scaling, worked the
best?
 In each case, where did bulblets form and how many were produced?
 If you were unsuccessful with a specific technique, what might be some reasons why you were
unsuccessful?
 If you were to try these propagation methods again, what might you do differently?
71
Asexual Propagation: T-Budding
[T; individual; 5 grafts/student]
Objectives:
 To better understand and demonstrate the techniques involved in T-budding.
 To successfully propagate a novelty plant using T-budding.
To Do Before Class:
Read pp. 23-25 in your lab manual and the sections on T-budding in your text (pp. 62, 114-115).
Plant Materials:
Hibiscus rosa-sinensis; named cultivars (Various selections of Chinese Hibiscus; Malvaceae, Mallow
Family) = scions (stock plants) & stocks (potted plants)
Materials To Bring To Class:




Pruners
Permanent marking pens
Polyethylene bags
Lab notebooks
Materials Provided:




Budding rubbers
Parafilm
Label stakes
Paper toweling
Methods:
1. Prepare the stock:
a. For this grafting exercise you will be using a plant that already has a root system as the stock.
Select a healthy, vigorous hibiscus plant from the assortment provided; the plants we will be
using are named cultivars and were propagated from softwood cuttings.
b. Starting near the base of the plant, and working up, you will be grafting five (5) buds, each from a
different hibiscus cultivar, onto the rootstock plant. Try and choose locations along the stem
72
that are relatively straight and evenly spaced along the lower portion of the stem.
c. Make the grafting cuts (See Figure 1). For each bud-graft, begin by making a 1 inch (2.5 cm)
long, vertical cut through the bark using a clean, sharp razor blade. Make a second, horizontal
cut centered on and just intersecting the top of the first cut to form a "T". Make sure your cuts
connect and go all the way through the bark (it is thicker than you think), yet not cut into the
underlying wood.
d. Using the corner of your razor blade, peel back the resulting flaps at the top of the "T" as if you
were opening a book, being careful to include the green cambium layer with the flap. Take care
to avoid contaminating the inner surfaces of the wound. The scion bud-piece will be inserted
into the resulting opening in the bark.
2. Prepare the scion:
a. When T-budding, the scion basically consists of a single bud with some attached bark and wood.
An assortment of hibiscus plants, again named cultivars, will serve as our source of budwood.
Choose the varieties you want to work with – you will need to collect five (5) buds; each from a
different variety than your rootstock plant. If all five buds are successful you will have created a
six-in-one hibiscus plant.
b. To collect the individual bud scions, choose healthy-looking, mature buds. Remove the leaf
blade from the subtending leaf, but leave the petiole intact to serve as a handle. To remove the
bud, undercut the bud starting your cut about 3/4 of an inch (1 cm) below the bud and
continuing up and beyond the bud for about 1 inch (2.5 cm).
3. Join the stock and scion to complete the graft:
a. Using the petiole as a handle, and the bud pointing up, insert the bud scion into the T-shaped
cut in the bark of the rootstock plant. Remove the excess bark above the bud by cutting it off
horizontally such that the scion fits flush under the edge of the bark at the top of the "T".
b. Starting below the graft, wrap the graft with a parafilm covering all the cut surfaces but not the
bud itself. Trim the petiole such that only a short stub remains.
c. Throughout the process, prevent excessive drying of the stock and bud scions. To this end,
work quickly and complete each graft separately. Buds may be temporarily wrapped in clean,
moist paper toweling in a polyethylene bag to prevent them from drying out.
4. Observe your grafts on a regular basis and record your observations. Avoid the temptation to
remove the parafilm to "sneak a peek" for at least several weeks. If the petiole remains green and
eventually abscises naturally the bud is probably in good shape. If, however, the petiole dries-up
and remains firmly attached, the graft has probably failed.
5. As always, be sure to clean up all work areas when you are finished.
73
Observations:

Observations should include (but not be limited to): callus formation, condition of the petiole
stub, appearance of the bud, bud break, and success or failure of your grafts.
Some Questions to Ponder:

Were your grafts successful or not? If not, Why not?

Why would plant propagators use T-budding?

What environmental conditions are important for successful T-budding?
74
Figure 1. T-Budding.
Preparing the stock
Preparing the scion
75
Joining the stock and scion
Asexual Propagation: Approach Grafting – Spudmatoes
[T; individual; 2 grafts/student]
Objectives:
 To better understand and demonstrate the techniques involved in approach grafting.
 To successfully propagate plants using approach grafting.
To Do Before Lab:
Read pp. 23-25 in your lab manual and p. 303 in your text.
Plant Materials:
Lycopersicon esculentum (Tomato; Solanaceae, Potato Family) = scion
Solanum tuberosum (Potato; Solanaceae, Potato Family) = stock
Materials To Bring To Class:
 Permanent marking pens
 Lab notebooks
Materials Provided:





Budding rubbers
Parafilm
Paper toweling
Bamboo stakes
Twist ties
Methods:
1. To create your spudmatoes, a grafted plant which will produce both potatoes and tomatoes, the
potato will need to serve as the rootstock since the potatoes are produced below ground. The
tomato plant will become the scion.
2. Select two (2) healthy potato plants and two (2) healthy tomato plants from the plug-grown plants
provided. Be sure you can tell them apart (note that the tomatoes have a distinctly different
smell!).
76
3. Repot the two plants as close as possible to each other in the center of one of the containers
provided.
4. Once planted, gently bring the stems of the two plants together to determine where the stems
naturally come into contact without undue force or contortion. Using a razor blade, remove any
leaves that are attached to the stems in this region.
5. Using a clean, sharp razor blade, carefully shave a thin (1/32 to 1/16 of an inch or about 1 mm
thick) strip from each stem in the region where the stems came in contact when pushed together in
Step 4. The wounded area should be 1 to 1.5 inches (2.5 to 4 cm) long. Be very careful; the
stems are delicate and it is very easy to cut right through them.
6. Push the stems together again such that the wounded areas are in solid contact. Carefully wrap the
graft with a strip of parafilm making sure the stems remain in solid contact (See Figure 1).
7. When you have finished wrapping the graft, stake the grafted plant using a bamboo stake and a
couple of twist ties.
8. Repeat the procedure with your remaining potato and tomato plants. Label your plants with your
name and the date.
9. Water your grafted plants and transfer them to the designated location in the greenhouse.
10. Observe your grafts on a regular basis and record your observations. Avoid removing the
wrapping to "sneak a peek" for at least five (5) weeks.
11. When the grafts have healed you can complete the process by severing the root system of the
tomato plant just below the graft union and the top of the potato plant just above the graft union.
Be careful to know for sure which stems you are cutting before you make your cuts. Keep the
plant staked to help reduce the chances of breaking the graft and use a razor blade to cut the
stems.
12. If the top of your grafted plant (tomato) does not wilt, it is a good sign that your graft has been
successful.
13. As always, be sure to clean up all work areas when you are finished.
Observations:
 Observations should include (but not be limited to): callus formation and success or failure of your
grafts.
Some Questions to Ponder:
 Were your grafts successful or not? If not, why not?
 What is the primary difference between the approach graft and the other grafts we have studied so
far? What, then, is the advantage of the approach graft?
77
Figure 1. Making an approach graft.
Preparing the stock and the scion
Joining the stock and the scion
78
Vegetative Propagation: Air Layering
[T; individual; 2 air layers/student]
Objectives:

To better understand and demonstrate the techniques involved in air layering.

To successfully propagate a house plant by air layering.
To Do Before Class:
Read the section on “Layering” in your lab manual (p. 22) and the section on air layering in your text
(pp.24-25 “Layering”, 64, and 105-107).
Plant Material:
Schefflera actinophylla (Umbrella Tree; Araliaceae, Aralin Family)
and/or Perhaps Others
Materials To Bring To Class:
 Permanent marking pens
 Polyethylene bags
 Lab notebooks
Materials Provided:




Label stakes
Bamboo stakes
Sphagnum moss (moistened)
Twist ties or string
Methods:
Go and get one (1) potted Schefflera plant.
1. Select a location approximately midway up the stem on each plant for your air-layers.
2. Carefully remove any shoots or leaves from the portion of the stem to be layered.
3. Wound the stem. This may be done by making a slanting cut upward into the stem or by girdling
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the stem. You may each of the plants in the same fashion or try a different technique with each
plant.
4. If a slanting cut is used, carefully use a razor blade to make the cut such that it extends into the
stem about 1/3 its diameter and is about 1 inch long. (Do not cut halfway into the stem or it could
easily break off.)
5. If girdling is chosen, girdle the stem by removing a ring of stem tissue about 1/4 inch (6 mm) wide
and 1/16 inch (2 mm) deep.
6. Cover the wounded area, and an additional inch or so on either side, with moistened sphagnum
moss. If the stem was wounded with a slanting cut, hold the cut open by inserting some of the
sphagnum into the open cut.
7. Cut open one of your polyethylene bags to create a flat sheet. Use this sheet of polyethylene film
to wrap the layer. The polyethylene wrapping should extend far enough beyond the moss to allow
it to be secured with string or twist ties.
8. Using label stakes, label your layers with your name and the date and place them in the designated
location in the greenhouse. Water your plants.
9. Observe your plants at least weekly and record your observations. If the sphagnum moss appears
to be drying out, carefully open the plastic sleeve and remoisten it. When you observe numerous
roots growing into the moss, remove the plastic sleeve and cut the stem below the root mass.
Carefully remove the excess moss and spread the roots, taking care to keep the roots moist at all
times. Plant the rooted layer in the planting media and pots provided. Water the new plant
thoroughly after potting.
10. As always, be sure to clean up all work areas when you are finished.
Observations:
 Observations should include (but not be limited to): success or failure.
Some Questions to Ponder:
 Where do new roots appear?
 Do you think using rooting promoters would improve success?
 If you were propagating your house plant on a commercial basis, how might you manipulate the
parent plant to increase propagation efficiency?
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Air-Layer
1.) Girdled Stem
2.) Peat Moss around girdled section
3.) Plastic fastened around Peat Moss and girdled area
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