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Hemp Descriptors v2 archived 2023-06-07

USDA Hemp Descriptor and Phenotyping Handbook, Version 2

ABOUT

Objectives

The USDA Hemp Descriptor and Phenotyping Handbook was undertaken with the following objectives:

To assist breeders and researchers in identifying accessions with specific traits to facilitate germplasm selection within hemp (Cannabis sativa L.) improvement programs.
To identify gaps in the existing hemp collections and help formulate strategies for future collection and conservation efforts.
To designate and maintain a core collection of critical materials.
To increase NPGS user utility and accessibility to hemp germplasm and associated data.
To identify duplicate accessions and reduce costs of hemp genetic resource conservation.

The methods and protocols are based on peer-reviewed literature and/or crowd-sourced from the hemp community. Robust, reliable, and high-dimensional data generated from these phenotyping efforts will empower conservation of hemp genetic diversity and aid selection of materials with unique trait combinations for breeding programs.

We have attempted to compile a list of standardized characterization and evaluation methods to capture passport information and to quantify morphology, horticultural and agronomic quality, pathogen resistance, and metabolic profile. This document can be used a reference to standardize phenotypic data collection across the broader pool of hemp germplasm and will be updated periodically as better methodologies emerge.

The information gained from these phenotyping efforts will be digitally stored and made publicly available within GRIN-Global alongside the hemp germplasm held within the Plant Genetic Resources Unit (PGRU) in Geneva, NY. An example of the germplasm that is held at PGRU can be seen here. Phenotypic summaries of PGRU hemp genetic resources can be accessed on GRIN here.

PGRU coordinates hemp germplasm collection and exchanges from domestic and foreign sources. Information related to plant genetic resources increases usefulness to diverse stakeholders. Phenotypic data can be collected either by the curator during routine multiplication or by collaborators during collection, germplasm screening, or breeding experiments. PGRU asks germplasm recipients or donors to provide as much data associated with these materials as possible.

Collected data can be stored in a spreadsheet using the trait_name as column headings and PUID as row names. Our lab prefers the conventions of “Tidy Data” 1,2. This document can then be emailed to the hemp germplasm curator (zachary.stansell@usda.gov) for inclusion into GRIN-Global. Please do not hesitate to reach out with any questions, comments, suggestions!

Versioning

This is version 2.0 of the USDA Hemp Descriptor and Phenotyping Handbook published in early 2023. Here, we try to draw on new experience, publications, conversations since the publication of version 1.0 in September 2021.

Small details are clarified throughout
Expanded sections:
- Pathology
- Agronomic evaluation
- Fiber quality
More protocols are added:
- Feral collection.
- Seed threshing
- Tissue culture
- Pollen collection
Additional citations are added

Editors

Zachary Stansell (Hemp Curator, USDA-ARS, Plant Genetic Resources Unit)
Tyler Gordon (Hemp Breeder, USDA-ARS, Plant Genetic Resources Unit)
Anthony Barraco (Biol Sci Technician, USDA-ARS, Plant Genetic Resources Unit)
Daniel Meyers (Biol Sci Technician, USDA-ARS, Plant Genetic Resources Unit)
[Anthony Rampulla] (Biol Sci Technician, USDA-ARS, Plant Genetic Resources Unit)
Anya Osatuke (Cornell Cooperative Extension)

The authors gratefully acknowledges the critical review, editing, and the numerous suggestions for improvements made by Ademola Aina, Olivia Aldin, Masoume Amirkhani, Anthony Barraco, Craig Beil, Gary Bergstrom, Mark Berhow, Peter Bretting, Charlie Brummer, Mark Bridgen, Kadie Britt, Korey Brownstein, Zachary Brym, Carlyn Buckler, Ali Cali, Brian Campbell, Craig Carlson, Jeffrey Carstens, Ernst Cebert, David Chalkley, Chengci Chen, Alyssa Collins, Whitney Cranshaw, Randy Crowl, Heather Darby, David Dierig, Jorge de Silva, Chris Delhom, Sadanand Dhekney, Shelby Ellison, David Fang, David Gang, Nicholas Genna, Heather Grab, Jason Griffin, Kelly Gude, Joshua Havill, Yu Jiang, Nick Kaczmar, Joanne Labate, Michael Loos, Jessica Lubell-Brand, Tyler Mark, Victoria Meakem, Virginia Moore, Maylin Murdock, Jay Noller, Luis Alberto Monserrate Oyola, Dániel Pap, Bear Reel, Andrew Ristvey, Moira Sheehan, Savanna Shelnutt, Chris Smart, Larry Smart, Faith Sparks, George Stack, Jeffrey Steiner, Conor Stephen, Alan Taylor, Jacob Toth, Daniela Vergara, and Don Viands.

The drawing on the front cover is used with permission by Anya Osatuke. Kadie Britt provided many primary source images and text for the invertebrate section. Craig Carlson provided original figures, methods, and many ideas. Jacob Toth, Joshua Havill, Savanna Shelnutt, Brian Campbell, Shelby Ellison, and Jeffrey Carstens provided many helpful comments, references, protocols, and edits. We have tried to acknowledge everyone who’s helped with this work, but any omissions are solely Zachary Stansell’s fault.

This work has drawn heavily on input from the Cornell Hemp Stakeholder Survey. Please take the survey if you have not already done so.

Please contact zachary.stansell@usda.gov with any questions, comments, remarks, or ideas.

The United States Department of Agriculture (USDA) prohibits discrimination in its programs on the basis of race, color, national origin, sex, religion, age, disability, political beliefs, and marital or familial status. Not all prohibited bases apply to all programs. Persons with disabilities who require alternative means for communication of program information (Braille, large print, audiotape, etc.) should contact the USDA Office of Communications at (202)720-5881 (voice) or (202)720-7808 (TOO). To file a complaint, write to the Secretary of Agriculture, U.S. Department of Agriculture, Washington, DC 0250, or call (202)720-7327 (voice) or (202)720- 1127 (TOO). USDA is an equal employment opportunity employer.

Boxes

Throughout this document there are special colored text boxes:

Phenotype/Descriptor

trait_name [datatype; units]

elevation_meters [decimal; m]
Elevation of collecting site above sea level.

🧪Phenotyping Protocol🧪

Seed germination

10 waterproof trays
Sterile water-holding material (cotton wool, paper towels)
200 seeds, stored between 4 to 56 weeks…

📐Equation📐

Percent moisture may be calculated as:

$\frac{(wet - dry)}{wet} \times 100\%$

📜List📜

Invertebrate pests

Acherontia atropos
Aculops cannabicolus
Aecidium cannabis
…

📚Additional References📚

One of the earliest publications lauding hemp was “The Praise Of Hemp Seed” by John Taylor -Taylor (1620).

Keywords

hemp
Cannabis sativa L.
germplasm
phenotype
trait
characterization
evaluation
USDA-ARS
NPGS
PGRU

Language

GRIN-Global supports displaying data in multiple languages for system-level data. That is, if the system requires text to be displayed that is not actual GRIN-Global data, that text should be in the appropriate language for the current user. This is accomplished by using a table ending with _lang as a child table.

Data types & units

Units
All units are SI unless otherwise indicated.

datetime
A datetime data type that can handle time in nanoseconds and has a year range extending from the year “0001” to “9999.”

decimal
The decimal data type can store a maximum of 38 digits, all of which can be to the right of the decimal point. The decimal data type stores an exact representation of the number; there is no approximation of the stored value.

int
The integer data type is stored as a 4-byte integer; numeric values can range from $-2^{31}$ through $2^{31} – 1$.

nvarchar
An nvarchar field can store a string of text characters (maximum 4,000). The “n” in nvarchar means uNicode. “varchar” is an abbreviation for variablelength character string. Essentially, nvarchar is variable text field that supports two-byte characters, therefore capable of handling non-English symbols.

PASSPORT

An accession consists of seed or plant material representing a sample of a single species, collected at a single time and location. An accession may be a sample of multiple plants found at the same location at the same time, or it may be collected from a single individual. By default, NPGS will retain different samples of a putative cultivar/population as discrete inventories nested within the Plant Introduction accession.

Accession

taxonomy_species_id [nvarchar]
Scientific name of accession linking the accession record to its taxonomy parent (genus / species). Modified from GRIN-Global. Subtaxon may be included:

‘subsp.’ (subspecies)
‘var.’ (variety; not the same as the breeder’s named variety [uniform & stable product of breeding] or cultivar.)
‘f.’ (form)
‘group’ (botanical variety not cultivar name )

PUID [nvarchar]
If persistent, unique identifier has been previously assigned, report. Assigned to one accession to be unambiguously referenced at the global level, with associated information aggregated via automated means. Genebanks not applying a true PUID should use a combination of Institute Code, Accession Number, and the Genus as a globally unique identifier. Modified from Bioversity International, FAO (2015).

improvement_status [nvarchar]

Short paragraph. If known, elaborate on material improvement status, e.g., wild, landrace, breeding material, hybrid, founder stock, colonal selection, mutant, polyploid, mapping population, transgenic, etc.

plant_name [nvarchar]

Top name assigned to display (sometimes referred to as the top name), typically given by farmer, breeder, seed-saver. Cultivar name is a possible type of top name. If in non-Latin alphabet, provide original spelling alongside a Latin-alphabet transliteration in remarks. Modified from GRIN-Global and Bioversity International, FAO (2015).

accession_pedigree [nvarchar]

Description of plant pedigree, if known, e.g.:

“Selection from ‘Carmagnola’”
‘Beniko’/‘Carmagnola’//‘Futura 75’///‘Carmaleonte’/‘Felina 32’//‘Futura 75’ * “Mutation found in ‘Beniko’”

ploidy [int]
Record ploidy if known. If mixoploid or other, elaborate in passport_remarks. See Adriel Garay and Sabry Elias (1998).

accession_ipr [nvarchar]
State PVP registration status, if applicable. U.S. Link.

Varieties may also be protected by a U.S. Plant Patent (e.g. CW2A) or Utility Patent and/or Plant Breeder’s Right from a UPOV country; e.g., Canada.

US patents search
UPOV requires account to search Variety Database.
Canada

crop_use [nvarchar]

Explain crop use(s); e.g., oil, fiber, secondary metabolite, ecosystem services.

Germplasm source

source_cooperator_id [nvarchar]
Field associating the cooperator (person or organization) who was the source of the germplasm. See Appendix.

collector_cooperator_id [nvarchar]
Indicating the individual collecting sample. See Appendix.

developer [nvarchar]
List the name of the organization (or person) that bred the material.

Sampling & location

Modified from S-1084 Collection Protocols, GRIN-Global, personal conversations with hemp researchers (Shelby Ellison and Jeffrey D. Carstens), and Bioversity International, FAO (2015) standards. See Appendix.

number_plants_sampled [int]
Number of plants sampled to collect the accession material ( S-1084 Collection Protocols). See Appendix.

source_date [datetime]
Date when germplasm is collected from source material ( S-1084 Collection Protocols).

geography_id [nvarchar]
The internal geographic identifier indicating the cooperator’s country and state ( S-1084 Collection Protocols).

elevation_meters [decimal; m]
Elevation of collecting site above sea level ( S-1084 Collection Protocols).

latitude & longitude [decimal]
Latitude and longitude in decimal degree format. The format is 10 integers and 8 decimals. Positive values are east of the Greenwich Meridian; negative values are west of the Greenwich Meridian ( S-1084 Collection Protocols).

coordinate_method [nvarchar]
Georeferencing method used (e.g.; GPS, map, estimated). Modified from Bioversity International, FAO (2015).

uncertainty [decimal; m]
Maximum coordinate uncertainty radius.

georeference_datum [nvarchar]
Geodetic datum/spatial reference system; WGS84 datum is preferred.

accession_inv_voucher_note [nvarchar]
If applicable, include additional voucher information.

ARCHITECTURE

Unless stated otherwise, measure plant architecture traits as the mean of 10 unpruned plants during week of sampling. Samples submitted to NPGS will be evaluated by a USDA-ARS laboratory using similar protocols as described below.

Morphology

Architectural traits modified from Carlson et al. (2021).

ht [decimal; cm]
Height of the stem from the ground to tip apical inflorescence, modified from (2021).

mcd [decimal; cm]
Maximum canopy diameter (mcd) as width of plant at widest set of branches (2021). Measured from widest tip to tip without stretching branches. Include flowering tissue in measurement.

mcdh [decimal; cm]
Height evaluated at maximum canopy diameter (mcdh) from ground to max canopy diameter (2021).

trkl [decimal; cm]
Trunk length (trkl) is evaluated as distance from ground to first branch (2021).

inl [decimal; cm]

Average internode length (inl) is calculated between internodes along the primary stem (50 cm max, see diagram) (2021).

$inl = \frac{ht-trkl}{branches}$

hyp_a and hyp_b [decimal; cm]

Calculated (2021).

$hyp.a = \sqrt{(ht - mcdh) ^ 2 + \frac{mcd}{2}^2}$.

$hyp.b = \sqrt{mcdh^2 + \frac{mcd}{2}^2}$

kite_perimeter [decimal; cm]

Calculated (2021).

$kite.perimeter = 2 \times (hyp.a + hyp.b)$

kite_area [decimal; $cm^2$]

Calculated (2021).

$kite.area = \frac{ht \times mcd}{2}$.

ba [decimal; (0-180°)]

Kite branch angle ba is calculated from the lower kite triangle, using the difference of maximum canopy diameter height and trunk length (2021).

$ba = \arctan \frac{mcd}{2(mcdh- trkl)}$

nodes | nodes_opp | nodes_alt [int]
Number of internodes (nodes) per plant; nodes are by definition $2 \times$ branches (2021). Number of opposite internodes (nodes_opp) per plant. Number of alternate internodes (nodes_alt) per plant. When grown from seed, branching is initially opposite, transitioning to alternate as the plant matures. Plants propagated from cuttings generally have alternate branching in the whole plant Stack et al. (2021).

branches [int]

Number of branches per plant (2021). When grown from seed, branching is initially opposite, transitioning to alternate as the plant matures. Plants propagated from cuttings generally have alternate branching in the whole plant Stack et al. (2021).

$branches = n.opp * (2 + n.alt)$

kite.circularity [categorical]

A continuous scale of apical dominance can be derived (2021):

$kite.circularity = \frac{4\pi\cdot kite.area}{kite.perimeter^2}$

dia [decimal; mm]
Diameter of the stem at soil level using calipers, forestry or fabric measuring tape, modified from Carlson et al. (2021).

pith_diameter [decimal; mm]
Diameter of the pith in the stem cross section at stem midpoint, modified from International Union for the Protection of New Varieties of Plants (2012).

Uncrewed aerial vehicle evaluation

uav_xxx [TBA]
See Carlson et al. (2021).

Remarks

architecture_remarks [nvarchar]
If possible, report date of measurement [days from sowing], sex average, minimum, and maximum height and width observed in a planting (cm).

📚Additional References📚

Anderson (1980)
Werf, Haasken, and Wijlhuizen (1994)
Werf et al. (1995)
Meijer and Keizer (1996)
Ranalli (1999)
Mishchenko and Lajko (2016)
Magagnini, Grassi, and Kotiranta (2018)
Backer et al. (2018)
Spitzer-Rimon et al. (2019)
Carlson et al. (2021)
Danziger and Bernstein (2021)
Stack et al. (2021)
Vergara et al. (2021)

LEAF

Unless otherwise noted, gather leaf data from the uppermost set of mature leaves, as mean of 5 leaves gathered from each of 10 different plants immediately before onset of flowering.

Morphology

petiole_length [decimal; cm]
central_leaflet_length [decimal; cm] central_leaflet_width [decimal; cm]

Leaf is flattened and measured from tip until start of rachis; petiole is flattened and measured from base of rachis until petiole base, modified from -International Union for the Protection of New Varieties of Plants (2012) and -Anderson (1980).

(A) central_leaflet_length and (B) petiole_length measurement.

Imaging

leaf_color_L [decimal]
leaf_color_a [decimal]
leaf_color_b [decimal]

The average color of uppermost set of mature leaves, collected before flowering, measured with a colorimeter, modified from -International Union for the Protection of New Varieties of Plants (2012). A RHS color chart may also be used, but values should be converted to (L*a*b*) before addition to GRIN. From Wikipedia (accessed 2023-01-11): The CIELAB color space, also referred to as L*a*b*, is a color space defined by the International Commission on Illumination […] in 1976. It expresses color as three values: L* for perceptual lightness and a* and b* for the four unique colors of human vision: red, green, blue and yellow. CIELAB was intended as a perceptually uniform space, where a given numerical change corresponds to a similar perceived change in color.

There are many programmatic solutions to convert colors (I use R for everything: 1,2,3) as well as many online tools (e.g., Colormine).

Consider printing a label to include in the scan as well. PGRU germplasm imaging and scans typically include accession ID, species, and plant id name (e.g. ‘FIN-314’). PGRU uses a small color wheel in the corner of our templates (DOWNLOAD), but that might not be necessary for you since you are measuring color with a colorimeter.

leaf_variegation [nvarchar; Y/N]

Indicate whether not variegation has been noted or is present.
Variegated leaves are most likely not virus based, but it might be worth investigating.

leaf_scan_protocol [.jpg or .png]

🧪Scan protocol🧪

Equipment

Flatbed scanner
Desktop monitor
Black cloth, ideally velvet, of equal dimensions to scanner bed.

Protocol

Gather one mature leaf from a representative sample of 10 plants a week before the onset of flowering. Retain petioles. Keep on ice if wilting is a concern.
Scan leaves within the hour of collection using a scanner with the lid open, draping black fabric over the leaves to absorb background light. Include scale or ruler (cm) and puid.
Convert the scanned leaf image into .png file and save.

Leaf imaging setup at PGRU

Leaf scan example from routine phenotyping at PGRU

📚Additional References📚

Haney and Kutscheid (1975)
Dayanandan and Kaufman (1976)
Anderson (1980)
Meijer and Keizer (1996)
Amaducci et al. (2008)
Hall, Bhattarai, and Midmore (2012)
Mishchenko and Lajko (2016)
Magagnini, Grassi, and Kotiranta (2018)
Spitzer-Rimon et al. (2019)
Carlson et al. (2021)
Stack et al. (2021)
Vergara et al. (2021)

SEX & INFLORESCENCE

General remarks

Phenotyping sex characters is (in our experience) more complex than it might seem initially. At PGRU, we have evaluated phenotypic sexual expression at both the plot level and by measuring many individual plants. Some groups have simply recorded the sex of 100 preselected individuals at harvest. Both approaches should produce the same phenotype. In our 2022 field trials in Geneva, NY, we recorded the exact first flowering date (of pistillate and staminate flowers) of > 2,400 individuals and then calculated (by population) the mean percent female, male, and monoecious. Specify your approach in sex_remarks, as well as sowing/transplanting date, field or greenhouse conditions, and photoperiod.

Hemp flowering guide produced by XXX with permission from

Sex ratio

sex_ratio [nvarchar]
Sex ratio during flowering.
Target a sample of 100 individuals in the field before flowering begins. Label and number sample plants clearly. Record F:M:O; as the ratio of female, male, and monoecious (individuals with male and female flowers) individuals, respectively.
If plants are wild-collected, 100 plants may not be available; measure as many as possible (See S-1084 Collection Protocols).

Phenology

date_flower_female [datetime]
date_flower_male [datetime]
date_flower_monoecious [datetime]

Pre-terminal date when axial flowers with shortening internodes and terminal pistils (clusters of flowers at shoot termini) were observed Carlson et al. (2021). Record sowing/transplanting date in sex_remarks. See also Faux et al. (2013); Shams et al. (2020).

days_2_female [integer; d]
days_2_male [integer; d]
days_2_monoecious [integer; d]

days_2_female = date_flower_female - sow date

days_2_male = date_flower_male - sow date

days_2_monoecious = date_flower_monoecious - sow date

maturity [int; d]
Days from germination to commercial maturity, calculated as harvest date - sowing date.

day_neutrality [nvarchar]
Describe flowering behavior as critical day length and/or day neutral response. Calculate as percent of plants in population exhibiting day neurtrality. A more precise definition is required to define this phenotype quantitatively.

Inflorescence

inflor_length [decimal; cm]
Length of inflorescence cluster at the uppermost branch, excluding leaves. Remove leaves with more than 1 leaflet. Retain stems and seeds; inflorescence may be retained as clusters.

inflor_weight [decimal; cm]
Measure weight after drying at room temperature for >48 hr.

inflor_yield [decimal; $kg\cdot ha^{-1}$]
Combine inflorescence dry weight data with planting density to calculate kg/ha.

inflor_color [nvarchar; L*a*b*]
Measured with a colorimeter at commercial maturity. A RHS color chart may also be used, but values should be converted to (L*a*b*) before addition to GRIN.

Remarks

sex_remarks [nvarchar]
Short paragraph, if possible/applicable include:

If plants were grown in field, greenhouse, or growth chamber.
Planting density (e.g., $\frac{seeds}{m^{2}}$) if applicable.
Female and male bract, stigma, and flower color.
Date of collection.
Day length (hh:mm) on the date of the first observed open female flower.

📚Additional References📚

Schaffner (1921)
Grishko, NN and Levchenko, VI and Seletski, VI (1937)
Werf, Haasken, and Wijlhuizen (1994)
Mandolino et al. (1999)
Ranalli (1999)
Lisson, Mendham, and Carberry (2000)
Shao, Song, and Clarke (2003)
Pahkala, Pahkala, and Syrjälä (2008)
Cosentino et al. (2012)
Hall, Bhattarai, and Midmore (2012)
Faux et al. (2013)
O. V. Razumova (2014)
Lynch et al. (2016)
Olga V. Razumova et al. (2016)
Vergara et al. (2016)
Punja, Rodriguez, and Chen (2017)
Zhang et al. (2018)
Eichhorn Bilodeau et al. (2019)
Salentijn, Petit, and Trindade (2019)
Kovalchuk et al. (2020)
Punja and Holmes (2020)
Toth et al. (2020)
Adal et al. (2021)
Danziger and Bernstein (2021)
Dowling, Melzer, and Schilling (2021)
Hurgobin et al. (2021)
Stack et al. (2021)

SEED

Samples submitted to NPGS will be evaluated by a USDA-ARS laboratory using similar protocols as described below.

General

hundred_seed_weight [decimal; g]
Record mass of 100 seeds.

thousand_seed_weight [decimal; g]
Record mass of 1000 seeds.

seed_image [.jpg or .png]

🧪SEED IMAGE🧪

Equipment

Flatbed scanner
Desktop monitor
Black cloth, ideally velvet, of equal dimensions to scanner bed.
Transparent, flat-bottomed tray or transparency film to protect scanner bed from scratches.

Protocol

Gather a sample of 20 grains.
Scatter samples across tray or transparency film.
Convert the scanned seed image into .PNG file and save.

Seed image. Photo credit Alan Taylor,Michael Loos, and Masoume Amirkhani

seed_size_length [nvarchar]
seed_size_width [nvarchar]

Size of the largest, smallest, and median seed in the lot (mm) as L:S:M using using Tomato Analyzer, Smart Grain, Photoshop, GNU Image Manipulation Program, or other imaging software to make these calculations based on a scanned image of 20 seeds.

seed_moisture [%]
Dry seeds in a single layer in a constant temperature oven held at 105 ˚C for 20 h.

$**`seed_moisture`** = \frac{(wet - dry)}{wet} \times 100\%$

Germination/viability

percent_germ [%]

🧪SPROUT TEST🧪

Equipment

10 waterproof trays
Sterile water-holding material (cotton wool, paper towels)
Water source
200 seeds, stored between 4 to 56 weeks after harvest in dry conditions between 16 - 25˚C.

Protocol

4 sub-samples of 50 seeds are isolated.
Each sub-sample is spread evenly on trays lined with water-holding material. Seeds may be placed on top of water holding material and in between rolls.
Each tray is saturated with water and placed in a temperature-controlled room: 16 hours of dark at 20˚C, 8 hours of light at 30˚C. Trays are kept moist.
Live, normal, sprouted seeds are counted by hand, sprouts are then removed from the germination tray.
After 7, 14, and 21, days the number of live, normal, sprouted seeds is counted by hand. The duration of experiment may be extended if dormancy is an issue (record in germ_remarks)
Germination percentage for each tray is calculated as: $\frac{ sprouted}{total} \times 100\%$ and averaged per tray.
Consider executing TZ tests for lots with high dormancy to verify they truly are dormant and not dead.

Seed germ testing (photo credit: Alan Taylor, Michael Loos, and Masoume Amirkhani).

Modified from Seed Laboratory Oregon State University (2018), NPGS protocols, and communication with Alan Taylor, Jeffrey Carstens, and Joshua Havill.

germ_remarks [nvarchar]
Record any dormancy issues and stratification methods used. Note: dormancy in feral/naturalized populations may also be important to characterize.
Cold stratification may be the norm in feral specimens with a temperate origin.

Seed threshing

See Appendix

Yield

grain_yield [decimal; kg/Ha]
Subsample larger strip trials (transect method) at seed maturity. Subsamples are chosen depending on the size of the field/plot to estimate kg/hectare.

shattering [nvarchar; L,M,H]
Retain subsample of field planting for shattering evaluation and report as low, medium, or high. A more precise definition is required to define this phenotype quantitatively. See Planteome ontology.

Oil

oil_content [decimal; %]
The percentage of oil constituting the whole, unhulled seed mass calculated as:

$\frac{extracted.oil}{ground.sample} \times 100\%$

Indicate whether measurement was made using Nuclear Magnetic Resonance (NMR) imaging, or using a Soxhlet extractor in seed_remarks.

🧪Soxhlet evaluation🧪

Seeds are ground to a particle size of 0.5 mm ± 0.1.
Place 10 g of ground seed into an extractor thimble, extracting with 100 mL hexane for 24 h at 70 ˚C.

If a different protocol is used, record the sample weight (g), solvent name (IUPAC) and volume (mL), extraction time (h) and extraction temperature (˚C).

Adapted from Devi and Khanam (2018).

🧪NMR evaluation🧪

To ensure accurate NMR evaluation of hemp seed, seed moisture content must be below 8%.
Subsample 10-15 seeds after evaluation of moisture content.
The sample is evaluated with a 40 mm diameter sample probe.
Prior to being placed in a sample box, seeds are to be further temperature-conditioned in the drying oven or a heating block at 40 ˚C for 90 min.
Instrument settings:
- 5 mHZ operating frequency
- 4 scans
- 1 second re-cycle delay
- 40 ˚C magnetic box temperature.
A calibration curve is constructed by comparing several varieties with a range of oil concentrations.
A linear calibration curve is constructed against the peak area of the NMR resonance signal normalized against sample mass.
The y-axis of the plot reports normalized peak area, and the x-axis reports % oil concentration.
Validate oil concentration from seeds of the same batch using Soxhlet extraction .
Indicate the % oil concentration of seed and the R2 value of the linear equation.
If a different NMR setting is used, list the weight of seeds sampled (g), seed temperature conditioning (˚C), operating frequency (mHZ) and line equation.

Adapted from Hutton, Garbow, and Hayes (1999). See also Shams et al. (2020); Yadav and Murthy (2016).

seed_fatty_acid

See ISO 12966-1:2014 gas chromatography protocols for determination of fatty acids, free and bound, in animal and vegetable fats and oils following their conversion to fatty acid methyl esters (FAMEs).

Protein

combustion_analysis [decimal]

🧪Protein combustion analysis🧪

150 mg seeds are frozen in liquid nitrogen and ground.
Seeds are placed into a combustion analyzer.
Combustion tube is at 960 °C, and oxygen is dosed for 80 s at 170 mL.

Adapted from Schultz et al. (2020).

🧪Kjeldahl method🧪

Based on an adaption of AOAC Official Method 991.20 by Thiex et al. (2002), where

Kjeldahl N % = ([std.acid sample (mL)] - [std.acid blank (mL)]) x ([HCl]) x 14.01) / (weight (g) x 10)

Crude protein % = % Kjeldahl N x 6.25

📚Additional References📚

Boyce (1900)
Mah (1923)
Tuma (1972)
Ribnicky et al. (2000)
Vaknin et al. (2011)
Sera et al. (2012)
Small and Brookes (2012)
Serkov (2015)
Suriyong et al. (2015)
Yadav and Murthy (2016)
Citti, Pacchetti, et al. (2018)
H. Hu, Liu, and Liu (2018)
Devi and Khanam (2018)
Jian et al. (2018)
Williams (2019)
He et al. (2020)
Moon et al. (2020)
Punja and Holmes (2020)
Schultz et al. (2020)
Serkov et al. (2020)

FIBER

Samples submitted to NPGS will be evaluated by a USDA-ARS laboratory using similar protocols as described below.

Yield

Hemp stem cross section. A = epidermis; B = bast and cortex; C = pith, also known as hurd, D = inner core, usually hollow but not at joints.

fiber_yield [decimal; $\frac{kg}{Ha}$]
Fiber yield; modified from Serkov (2015). See also Backer et al. (2018). Harvest 10 plants, remove top and bottom 15 cm, defoliate run sample through a chipper, and measure mass. Calculate fiber_yield based on planting density.

wet_fiber_biomass [decimal; g]
Harvest stems 15 cm from base, remove top 15 cm, and defoliate. Wet biomass is average weight per stem without inflorescences from a sample of 10 plants Modified from Carlson et al. (2021).

dry_fiber_biomass [decimal; g]
Use sample from wet_fiber_biomass measurements. Dry biomass is average weight per stem dried at 65 ˚C until brittle from a sample of 10 plants. Modified from Carlson et al. (2021).

bast [decimal; %]
Stems collected from a sample of 10 plants, dried until brittle, and separated into bast (bark) and hurd (core) using a flax breaker, or by hand. Record as fraction of oven-dried mass.

hurd [decimal; %]
Stems collected from a sample of 10 plants, dried until brittle, and separated into bast (bark) and hurd (core) using a flax breaker, or by hand. Record as fraction of oven-dried mass.

📐Bast & hurd content📐

bast = $\frac{bast}{dry} \times 100\%$

hurd = $100\% - bast$

bast_soluble & fiber_solubility [decimal; %]
Soluble materials in bark after evaluation with sodium hydroxide evaluation.

🧪Sodium hydroxide evaluation🧪

Protocol

10 - 15 g isolated hurd samples isolated from dry stems, or 10 - 15 g isolated bast samples from dry stems.
Pulp dispersion apparatus, consisting of a variable speed motor and a stainless steel stirrer with a shell.
Thermometer
Vacuum source
Filtering flasks, 100 mL
Graduated cylinder, 100 mL
Watch glasses
Stirring rods
Tall, 200 mL beakers
Water bath with cover and holes to securely submerge the bottoms of tall, 200 mL beakers.
Filtering crucibles, 30 mL, 10 - 15 µm maximum pore size.
1000 mL sodium hydroxide solution, 1.0 % ± 0.1 NaOH (0.25 N)
1000 mL acetic acid, 10%.

Grind hurd or bast sample into a fine meal.
Dry filtering crucibles before use in an oven at 105° ± 3 °C for 60 min, cool in a desiccator, and record weight.
Heat water bath to 97 - 100 ˚C
Adjust speed of the motor and the angle of the blades so that no air is drawn into the pulp suspension during stirring.
Place 10 g meal into 200 mL beaker and add 100 mL 1% NaOH and stir with glass rod.
Cover beaker with watch glass and place into water bath for 1 h. Keep water level above level of alkali solution in the beaker at all times.
Stir the meal with a rod for about 5 s at 10, 15, and 25 min after placing into the bath.
At the end of 1 h, transfer the material to a tared filtering crucible and wash with 100 mL of hot water.
Add 25 mL of 10 /% acetic acid and allow to soak for 1 min before removal. Repeat this step once again.
Wash material with 100 mL hot water three times to remove acid.
Dry crucible and contents in the oven at 105 - 108 ˚C until constant weight.
Cool in desiccator.

📐% Soluble materials📐

$\frac{meal.before - meal.after}{meal.before} \times 100\%$

non_fiber [TBD]
Protocol for evaluating stem non-fiber components not determined.

Quality

There is little modern research on hemp fiber quality evaluation protocols. This work will be conducted in collaboration with the Southern Regional Research Center (SRRC).

📜Candidate fiber traits📜

fiber length
fiber strength
fiber flexibility; see Werf, Haasken, and Wijlhuizen (1994)
fiber length/diameter ratio; see Ranalli (1999)
tensile strength; see Ranalli (1999)
brittleness; see Ranalli (1999)
crystallization/cellulose crystallinity Rongpipi et al. (2019)
cross linking
elasticity; see Ranalli (1999)
ease of decortication
mechanical vs microbial retting
cellulose:lignin ratio

fiber_remarks [nvarchar]

Remarks are used to add notes or to elaborate on fiber descriptors

📚Additional References📚

Charles (1708)
Unknown (1720)
David (1729)
Slator (1735)
Marcandier (1764)
Gee (1767)
Farmer (1775)
Wissett (1808)
Humphrey (1919)
Vavilov (1957)
Werf et al. (1995)
Daryl T. Ehrensing (1998)
Rustichelli et al. (1998)
Hampton (2000)
Struik et al. (1999)
Hillig (2005)
Pahkala, Pahkala, and Syrjälä (2008)
Zatta, Monti, and Venturi (2012)
Piluzza et al. (2013)
Salentijn et al. (2015)
Serkov (2015)
Mishchenko and Lajko (2016)
Grassi and McPartland (2017)
Tang et al. (2016)
Weijde et al. (2016)
Johnson (2018)
Musio, Müssig, and Amaducci (2018)
Salentijn, Petit, and Trindade (2019)
Williams (2019)
Petit et al. (2020)

SECONDARY METABOLITES

Unless otherwise stated, collect from a sample of 10 plants at harvest. Samples submitted to NPGS will be evaluated by a USDA-ARS laboratory using similar protocols as described below.

Chemotype

chemotype [int; 1-6]
Note that living tissues synthesize acid forms of most cannabinoids (i.e. THC-a, CBD-a) and these are often decarboxylated to non-acid forms during evaluation (i.e. THC, CBD). Chemotype is largely driven by segregation of alleles at the $B$ or $O$ loci Toth et al. (2020).

1 = Mostly THC(A)
2 = ~1.5:1 CBD(A):THC(A)
3 = Mostly CBD(A)
4 = Mostly CBG(A)
5 = Low overall cannabinoid content
6 = Other

chemotype_segregation [nvarchar]

To not mask chemotype, we recommend measuring cannabinoids from 10 individual plants rather than pooling samples. If multiple individuals are evaluated for chemotype and differing chemotypes are observed in the same population, indicate the percentage of each chemotype using the chemotype scale (1:#;2:#;3:#,4:#,5:#).

Cannabinoids

cannabinoids [decimal; $\frac{μg}{mg}$]
Cannabinoid content is evaluated either by UPLC or HPLC. THC is a ‘’sticky’’ compound and residues will persist on laboratory equipment and analytical vessels following cleaning. THC carryover onto subsequent samples may erroneously increase the reported THC content. We strongly advise against re-using sample vials for UPLC and HPLC cannabinoid evaluation.

UPLC uses a shorter run time per sample, however HPLC provides better resolution for minor cannabinoids. Both methods are sufficient for quantifying total THC in compliance with the 2021 USDA-AMS Hemp Final Rule.
10 samples collected from 10 individual plants. Each sample should be analyzed in triplicate. Cannabinoid content is variable across the height of the plant. THC-a decarboxylates into THC following exposure to heat. We advise against decarboxylating cannabis samples prior to instrument evaluation, as this process introduces error through the volatilization of a variable percentage of the total cannabinoid content.

Instead, we recommend combining the THC-a and THC content of unheated plant samples to calculate the total THC content. The formula for this is:

📐THC & Cannabinoid analytes📐

$THC_{total.potential}(μg/mg) = THC + 0.877 \cdot THC_a$

$Cannabinoid.analyte = C_e\frac{V_f}{W_s}$

where:

$C_e$: Conc of the sample in the extraction solution ($\frac{μg}{μL}$)
$V_f$: Final volume of the sample ($μl$)
$W_s$: Weight of the sample ($mg$)

🧪UPLC cannabinoid evaluation🧪

Sample run time: 10 min.

Equipment

Sonicator
Thermo Scientific UltiMate 3000 controlled by Chromeleon software
IntertSustain C18 3 μm chromatographic column (2.1 x 100 mm)
5 mM aqueous ammonium formate
0.1% formic acid
95% formic acid
Acetonitrile
Volumetric flasks: 1.5 L, 50 mL.
0.45 µm membrane filter.

Standards:

Cannabidivarinic Acid (CBDVA; CAS #: 31932-13-5)
Cannabigerovarinic Acid (CBGVA; CAS #: 64924-07-8)
Cannabidivarin (CBDV; CAS #: 24274-48-4)
Cannabivarin (CBV; CAS #: 33745-21-0)
Cannabidiolic Acid (CBDA; CAS #: 1244-58-2)
Cannabigerolic Acid (CBGA; CAS #: 25555-57-1)
Tetrahydrocannabivarin (THCV; CAS #: 31262-37-0)
Cannabigerol (CBG; CAS #: 25654-31-3)
Cannabidiol (CBD; CAS #: 13956-29-1)
Tetrahydrocannabivarinic Acid (THCVA; CAS #: 39986-26-0)
Cannabichromevarinic Acid (CBCVA; CAS #: 1628112-69-5)
Cannabinol (CBN; CAS #: 521-35-7)
Cannabinolic Acid (CBNA; CAS #: 2808-39-1)
Δ9-Tetrahydrocannabinol (THCd9; CAS #: 1972-08-3)
Δ8-Tetrahydrocannabinol (THCd8; CAS #: 5957-75-5)
Δ9-Tetrahydrocannabinolic acid A (THCA or THCAA; CAS #: 23978-85-0)
Cannabicyclol (CBL; CAS #: 21366-63-2)
Cannabichromene (CBC; CAS #: 20675-51-8)
Cannabichromenic Acid (CBCA; CAS #: 185505-15-1)
Cannabicyclolic Acid (CBLA; CAS #: 40524-99-0)

Standard preparation

All standard solutions are to be prepared at a concentration of 100 µg/mL.
Cannabinoid acid standards are prepared in acetonitrile. Decarboxylated cannabinoids are prepared in methanol.

Reagent preparation

1 L reagent for Mobile Phase A: Combine 0.21 mL 95% formic acid with 5 mM ammonium hydroxide (0.62 mL 30% ammonium hydroxide solution) with millipore water to 999 mL. Add 1 mL formic acid.
1 L reagent for Mobile Phase B: Combine acetonitrile with formic acid to 0.1% formic acid.

Protocol

Dry inflorescences at room temperature (20 °C) to < 10% moisture.
Grind samples.
Sieve through a number 20 sieve (850 μm).
Place 0.100 grams of ground hemp material in a 20 mL glass scintillation vial.
Prepare each sample in triplicate.
Calibrate the pipetter to ensure it is delivering the correct volume of 200-proof ethanol. Check that the weight of 20 mL of ethanol delivered is 15.78 g (density: 0.789 g/cm³).
Add 20 mL of HPLC grade 200-proof ethanol to the sample.
Sonicate (40 kHz) in a sonication bath for 30 min at room temperature.
Remove from sonication bath.
Allow the sample to sit overnight (18 h) in the dark at room temperature.
The next day filter the supernatant through a 0.45 μm Nylon membrane filter into a sample vial for analysis.
Inject 1 µL sample volume.
Set flow rate to 0.60 mL/min.
UPLC gradient program: 0-2 min, 70% B (concave curve 9); 2-5.5 min, 75% B (concave curve 9); 5.5-6 min, 100% B (linear curve 5); 6-7 min, 100% B; 7-7.5 min, 65% B (linear curve 5); and 7.5-8, 65% B followed by 2.0 min of equilibration at 65% B between injections.
The autosampler chamber and column temperature are set to 8°C (if a temperature-controlled sample tray is available) and 50°C, respectively. .
Five-point standard curves are developed for the cannabinoids using a setting of 228 nm from the diode array detector.

Methods provided by Berhow and Gude (2021) and modified by Dr. Korey Brownstein in 2023.

Other metabolites

terpenes [decimal]

🧪Terpene evaluation🧪

Equipment

GC/MS (Agilent 7890B GC, Agilent 5977B MSD, PAL 3) controlled by LabSolutions software
HP-5MS UI, 30 m × 0.25 mm, film 0.25 μm column
0.45 μm filter
Hexane

Standards

β-bisabolol
Bulnesol
m-Camphorene
p-Camphorene
Δ3-Carene
β-Caryophyllene
10-epi-γ-Eudesmol
α-eudesmol
β-eudesmol
α-humulene
Limonene
Linalool
β-Myrcene
Plastochromanol-8
α-Phellandrene
α-Pinene
cis-Sabinene hydrate
γ-Selmene
Selna-3,7(11)-diene
α-Tocopherol
β-Tocopherol
δ-Tocopherol
γ-Tocopherol
β-Pinene
Nerolidol
Camphene
Terpinolene
Ocimene
α-Terpinene
γ-Terpinene

Protocol

Dilute essential oil standards with hexane to a concentration of 0.1 mg/mL.
Dry inflorescences at room temperature (20 ˚C) to > 10% moisture.
Grind samples.
Combine 20 mg plant material with hexane.
Pass through the 0.45 µm filter to extract terpenes.
Dilute filtrate 1:20 with hexane.
Inject samples into split injection port (1:5).
Instrument settings as follows:
- Injection port held at 100 ˚C with an initial time of 4 min.
- Inlet temperature fixed at 250 ˚C.
- Detector temperature fixed at 280 ˚C.
- Column held at 35 °C for 5 min, then raise to 150 °C at 5 °C/min. Then raise to 250 ˚C at 15˚ C/min. Hold time 90 min.
- Helium serves as carrier gas, with flow rate 1 mL/min.

Adapted from Hanuš and Hod (2020).

Remarks

metabolite_remarks [nvarchar]

Identification of other metabolites may be provided in the metabolite_remarks field.

📚Additional References📚

Turner, Hemphill, and Mahlberg (1978)
Rustichelli et al. (1998)
Meijer and Hammond (2005)
De Backer et al. (2009)
USDA (2009)
Casano et al. (2011)
Russo (2011)
Hazekamp and Fischedick (2012)
Pertwee (2014)
Pandohee et al. (2015)
Lynch et al. (2016)
Weijde et al. (2016)
Brighenti et al. (2017)
Dufresnes et al. (2017)
Jin et al. (2017)
Patel, Wene, and Fan (2017)
Ciolino, Ranieri, and Taylor (2018)
Citti, Braghiroli, et al. (2018)
Citti, Pacchetti, et al. (2018)
Palmieri et al. (2018)
Pellati et al. (2018)
Pollastro, Minassi, and Fresu (2018)
Wang et al. (2018)
Zivovinovic et al. (2018)
Booth and Bohlmann (2018)
Comeau et al. (2018)
Hädener, König, and Weinmann (2018)
Laverty et al. (2018)
Mandrioli et al. (2018)
Protti et al. (2018)
Reimann-Philipp et al. (2018)
Pavlovic et al. (2019)
Nahar, Onder, and Sarker (2020)
Križman (2020)
Danziger and Bernstein (2021)
Hurgobin et al. (2021)
Stack et al. (2021)

PATHOGEN/PEST

Diseases are scored by indicating% area of planting affected and whether crop failure occurred in conjunction with infection. If one data score is provided for multiple plantings, provide the highest area coverage observed. Ideally the hemp community should decide on a set of check-varieties to include in every trial to compare between seasons.

For consistent scoring of disease and stress data, it is recommended that the same individual be responsible for rating the entire planting.
Refrain from capturing data for a small set or singular accessions, instead prefer an actual growout of a replicated trial that contains numerous accessions in a single season.

Record other relevant information in the disease_remarks field. Specifically, it will be useful to distinguish between hop powdery mildew, caused by P. macularis, and cannabis powdery mildew, caused by Golovinomyces ambrosiae. These appear to possess varying degrees of aggressiveness and phenotyping remarks should include information about which pathogen species was used for phenotyping.

Fungal and oomycetes

Individual plants are visually inspected for powdery mildew severity based on a continuous scale of 0–100% plant area showing disease symptoms according to Stack et al. (2021). Three powdery mildew severity examples are shown below.

Powdery mildew on hemp; photo credit Tyler Gordon

fungal_xxx [decimal; %]
Known or suspected fungal/oomycete pathogens:

Alternaria spp
Ascochyta spp
Athelia rolfsii or Sclerotium rolfsii
Bipolaris spp
Bipolaris gigantea
Boeremia
Botrytis cinerea
Botrytis pseudocinerea
Botrytis porri
Cercospora cf. flagellaris
Chaetomium globosum
Cladosporium spp
Colletotrichum spp
Curvularia spp
Diaporthe eres/ D. subordinaria
Exserohilum spp
Fusarium avenaceum
Fusarium brachygibbosum
Fusarium chlamydosporum
Fusarium equiseti
Fusarium graminearum
Fusarium lichenicola
Fusarium oxysporum
Fusarium proliferatum
Fusarium solani
Fusarium sporotrichiodes
Fusarium tricinctum
Globisporangium irregulare
Globisporangium ultimum
Golovinomyces ambrosiae
Golovinomyces cichoracearum
Golovinomyces spadiceus
Lasiodiplodia theobromae
Leveillula taurica
Leptosphaeria
Neofusicoccum parvum
Penicillium spp
Phoma spp
Phoma multirostrata
Phomopsis spp
Phytophthora spp
Podosphaera macularis (syn. Sphaerotheca macularis)
Pseudocercospora spp
Pythium aphanidermatum
Pythium catenulatum
Pythium dissotocum
Pythium myriotylum
Rhizoctonia solani
Sclerotinia sclerotiorum
Sclerotinia minor
Sclerotium rolfsii
Septoria spp
Stemphylium spp
Thielaviopsis basicola
Uredo kriegeriana
Verticillium dahliae

Bacterial

bacterial_xxx [decimal; %]
Known or suspected bacterial pathogens:

Agrobacterium tumefaciens
Pseudomonas koreensis
Pseudomonas syringae
Serratia marcescens
Sphingomonas yanoikuyae
Xanthomonas campestris pv. cannabis

Virus/Viroid

virus_xxx [decimal; %]
Known or suspected virus/viroid/phytoplasma pathogens:

Alfalfa mosaic virus (AMV)
Arabis mosaic virus (ArMV)
Beet curly top virus
Cannabis sativa mitovirus 1
Cannabis cryptic virus
Citrus yellow-vein associated virus
Cucumber mosaic virus (CMV)
Curly top virus
Lettuce chlorosis virus (LCV)
Tobacco ringspot virus (TRSV)
Tomato ringspot virus (ToRSV)
Tobacco streak virus (TSV)
Tomato mosaic virus (ToMV)

viroid_xxx [decimal; %]

Hop latent viroid (HLVd)

phytoplasma_xxx

Candidatus phytoplasma trifolii

nematode_xxx

Meloidogyne incognita (Root knot nematodes)

Remarks

disease_remarks [nvarchar]

In a short paragraph, describe the disease. Please include the anatomy of the plant affected, the growth stage of the plant affected, symptoms of infection, patterns of spread in field or greenhouse, evidence leading to conclusion of the nature of the pathogen, and any other relevant observations.

Invertebrate

Key pests
Many insects and mites can be observed in hemp and the pest complex can differ depending on whether the crop is cultivated indoors or outdoors. While many arthropods can be found in hemp, some of the most often-seen pests include corn earworm, twospotted spider mite, cannabis aphid, and hemp russet mite.

Invertebrate pests are scored by indicating % area of planting affected and whether crop failure occurred in conjunction with infestation.

Refrain from capturing data for a small set or singular accessions, instead prefer an actual growout of a replicated trial that contains numerous accessions in a single season. Ideally the hemp community should decide on a set of check-varieties to include in every trial to compare between seasons.

Images and text were very generously provided by Kadie Britt -Kadie Britt (2021). We highly recommend John Michael McPartland, Clarke, and Watson (2000), W. Cranshaw et al. (2019), and Britt (2021) as excellent overviews.

Corn earworm

helicoverpa_zea [decimal; %]
Helicoverpa zea or corn earworm is the most damaging pest of hemp grown in outdoor environments as it targets marketable portions of hemp plants – floral regions of CBD and seeds of grain varieties -Kadie Britt (2021).

Young corn earworm larva. (Photo credit: Kadie Britt)

Bud rot in floral hemp bud from corn earworm feeding. (Photo credit: Kadie Britt

Adult corn earworm moth on hemp plant. (Photo credit: Katlyn Catron)

Hemp russet mite

aculops_cannabicola [decimal; %]
Aculops cannabicola or hemp russet mite is a microscopic, cannabis-specific mite that can be found in indoor and outdoor hemp. Mites have four legs on their white- to beige-colored, cigar shaped bodies and are not visible without the use of magnification Kadie Britt (2021) and John M. McPartland and Hillig (2003).

Hemp russet mites on underside of hemp leaf. (Photo credit: Kadie Britt)

Upward curling of hemp leaves. Can sometimes be a symptom of hemp russet mite feeding injury to hemp but depends on cultivar. (Photo credit: Kadie Britt)

Twospotted spider mite

tetranychus_urticae [decimal; %]
Tetranychus urticae or twospotted spider mite is a generalist mite pest that can be found indoors and outdoors. Feeding injury causes stippling marks on leaves (Figure 1.7) and webbing can sometimes be observed in apical portions of plants (Figure 1.8). This mite is small and oval in shape, has 8 legs, and can be orange/red or brown with two distinct dark spots on the body -Kadie Britt (2021).

Twospotted spider mite on hemp leaf. (Photo Credit: Kadie Britt)

Stippling on leaves due to twospotted spider mite feeding. (Photo credit: Kadie Britt)

Webbing in apical portion of plant due to twospotted spider mite populations. (Photo credit: Kadie Britt)

Cannabis aphid

phorodon_cannabis [decimal; %] Phorodon cannabis or cannabis aphid is a specialist, piercing-sucking insect that feeds exclusively on hemp. Populations can rapidly increase in favorable environments (Figure 1.10) as aphids can reproduce via asexual reproduction Kadie Britt (2021) and W. S. Cranshaw et al. (2018).

Cannabis aphid infestation on indoor hemp plant. (Photo credit: Kadie Britt)

Cannabis aphid skins caught in honeydew on surface of hemp leaves. (Photo credit: Kadie Britt)

Other pests

Known or suspected pests:

Acherontia atropos
Aculops cannabicola
Aecidium cannabis
Agallia constricta
Aglais urticae
Agromyza reptans
Aphis fabae
Aphis gossypii
Camnula pellucida
Ceutorhynchus assimilis
Chinavia hilaris
Chloealtis conspersa
Chlorochroa ligata
Chlorochroa uhleri
Chromatomyia horticola
Cosmopepla lintneriana
Bemisia tabaci
Diabrotica undecimpunctata howardi
Ditylenchus dipsaci
Empoasca fabae
Estigmene acrea
Euschistus servus
Frankliniella fusca
Frankliniella occidentalis
Graphocephala versuta
Grapholita delineana
Halyomorpha halys
Helicoverpa zea
Heterodera humuli
Hysteroneura setariae
Liorhyssus hyalinus
Liriomyza cannabis
Liriomyza strigata
Loxostege sticticalis
Mamestra configurata
Melanchra picta
Melanoplus bivittatus
Melanoplus femurrubrum
Melanoplus lakinus
Melanoplus differentialis
Microtechnites bractatus
Micrutalis calva
Miridae
Nezara viridula
Oebalus pugnax
Ostrinia nubilalis
Pentatomidae
Peridroma saucia
Phorodon cannabis
Phyllophaga tristis
Phyllotreta pusilla
Podosphaera macularis
Polyphagotarsonemus latus
Popillia japonica
Prionus
Pseudoperonospora cannabina
Pseudoperonospora humuli
Psylliodes attenuata
Rhopalidae
Rhopalosiphum abdominalis
Spilosoma virginica
Spissistilus festinus
Spodoptera exigua
Spodoptera ornithigalli
Strymon melinus
Systena blanda
Systena elongata
Tetramorium caespitum
Tetranychus urticae
Thamnurgus caucasicus
Thrips tabaci
Thyanta custator
Trichiocampus cannabis
Uredo kriegeriana
Vanessa cardui
Known European species

📚Additional References📚

H M G Van der Werf, W C A van Geel, and M Wijlhuizen (1995)
Punja, Rodriguez, and Chen (2017)
W. Cranshaw et al. (2019)
Eric Anderson (2019)
Campbell et al. (2019)
McKernan et al. (2020)
Szarka et al. (2020)
Farinas and Peduto (2020)
Thiessen et al. (2020)
J. Hu, Masson, and Dickey (2020)
Stack et al. (2021)

APPENDICES

Feral Hemp Collection

Feral hemp collection protocol by Dr. Shelby Ellison and Dr. Ademola Aina at University of Wisconsin Madison. Germplasm collected from this work COLLECTION, PRE-CHARACTERIZATION, AND PRESERVATION OF AMERICAN HEMP GERMPLASM will be donated to the Plant Genetic Resources Unit.

🧪Feral Hemp Collection🧪

Materials needed:

Paper bags #10 (6.5 x 4 x 13.25”) (50 per population)
Reflective vests
Waterproof folders/bags for datasheets
Datasheets
Sharpies
Pens

Tools required:

Gloves
Tape measure
Camera (Phone’s camera is okay)
Loppers
Pruning shears

Protocol

Secure state or federal licensing to possess industrial hemp in your jurisdiction.
Secure written authorization from your regulatory agency (state department of agriculture) and the University or institutional legal counsel (if required) to collect and hold feral hemp that has not been certified as < 0.3% $9-THC. Maintain copies of permits, licenses, and/or written authorization during fieldwork and with collected material through all subsequent steps in the protocol.
Investigate locations identified by citizen scientists (either through iNaturalist or word of mouth) as potential feral hemp populations. Avoid cultivated fields of industrial hemp (with potential proprietary genetics) and cultivated marijuana, be it state-registered or clandestine and illegal. Endeavor to cover the broadest range of ecoregions that can feasibly be accessed during fieldwork.
Secure verbal and/or written permission from the landowner/tenants to collect plant material.
Collect GPS coordinates and take pictures of the plants while capturing their immediate surroundings using a smartphone. Upload new coordinates along with pictures on iNaturalist for future collection purposes. Record a brief description of the site – drainage patterns, surrounding vegetation, elevation, and soil classification from the web soil survey on the provided datasheet.
1. Preferred population size > 50 female individuals, at a minimum of 10 females.
2. Populations separated by at least 5 miles (8km) are preferred to increase the likelihood of pollen isolation.
3. Give each population an ID code in this scheme: state (WI), year (23), collector’s initials (SE), two-digit sequential program number (01), and plant number (01) -i.e. WI-23-SE-01-01.
4. Upload photos to a shared computer drive for backup
Locate up to 50 female (seed-bearing) plants. For each plant, label a large paper bag with an ID code. Chop down at the base of plant with the loppers, lay on the ground, and measure length (primary meristem to base of plant) with a tape measure, record the ID code and length in the nearest inch on the datasheet. Use the pruning shears to cut the terminal and lateral seed-bearing branches into 6-8” sections and fill the bag up to ¾ of its height, allowing enough space to fold the top of the bag over. Do not collect more material than would fit a single bag. Collecting each plant in a single, separate bag is essential. If there are more seed-bearing branches on a plant than fit the bag, you might prioritize the most heavily seeded branches but do not overstuff the bag and do not put material from multiple plants in the same bag.
Hot, moist conditions are detrimental to seed viability. Bagged material should begin drying within 12 hours of collection. At no point should the paper bags be enclosed in plastic. Samples should be kept as cool as possible during transport from the field to a location where they can be dried.
Dry the bags at room temperature with a fan to move air for at least one week. Seed and plant material can be placed in a drying oven as long as temperatures are less than 35 °C (95 °F). Conditions exceeding 35 °C (95 °F) are detrimental to viability.
Once the plants are dried, hand-strip the branches to remove the bulkier stems from each bag. Stems may be discarded in compost or as waste. Fold the top of the bag over and staple or tape it closed.
Line in a shipping box with a plastic bag, place paper bags and the datasheet(s) inside the plastic bag, seal the box. Attach the shipping label provided and arrange for a Fedex pickup or deliver to a Fedex drop-off location. Refer to the contents only as “Dried plant material for scientific research”. Not hazardous and of no commercial value.

Data collection sheet should have the following details:

Collector’s name
Date (YYYY-MM-DD)
Location (verbal)
Latitude and Longitude (decimal)
State
Population #
Site Description
Column Headers: Plant, Length (cm)

Threshing

🧪Threshing🧪

Hemp has valuable seed and grain that can be difficult to harvest by hand, especially in large quantities.

Example sample from PGRU trials

PGRU runs fatty acid and protein analysis for all germplasm held in the repository using three small, electric Almaco BT 14 belt threshers.

Belt threshing rig at PGRU

To safely and efficiently remove and separate seed from hemp stalks, PGRU follows the following protocol:

Personal Protective Equipment (PPE)

Protective eye wear, ear plugs, N-95 mask, gloves, closed-toed shoes, long pants, long sleeve shirt.

Useful Items

Trash bin, pruning shears, working table, bins for seed collection and chaff, vacuum, air compressor.

Procedure

Verify the room you are using is well ventilated, turn on system.
Verify the machine was cleaned after last use. Ensure that the thresher has been properly maintained and the belts, chain, and electric connections are in good working condition.
Move thresher into open space and let others know that you will be threshing (generating dust and noise).
Set up a seed collection tray on the back end of the thresher under the seed collection compartment.
Turn machine on and verify both belts are operating smoothly. The belt speed can’t be adjusted. The distance between belts can be adjusted.
The fan on the thresher has variable air flow and can be turned off depending on the application.
Set up a line of bags and a trash bin on the side you will be threshing.
Pull individual stalks from the harvest bag and inspect them for seeds/chaff, if no seed or chaff is observed, discard the stalks. If seed or chaff is observed, hold the base of the stalk, and place it on the tray of the thresher. Slide the stalk into the running belts of the thresher and let the stalk go through the thresher. Sometimes the thresher is unable to remove all the seed, if this happens manually remove the seeds from the stalk onto belt. When all the seed has been removed, discard the stalks. Continue this until all stalks have been removed from the bag.
The harvest bag will have smaller stems, seed, and chaff after the larger stalks have been removed. Empty the remaining bag contents into the thresher.
Remove any large stalks from the seed collection bin and empty the collected seeds and chaff back into the harvest bag.
Roll or fold the bag down to a small size and place the bag in a cool dry room for further processing.
Turn off the thresher and clean the area at the end of the workday.

Considerations

Consider the max diameter of half an inch for stalks that can go through the belts
Constant monitoring of your airflow can be achieved by checking for clean seed (lacks chaff and stalk)
One full sized stalk should only take 10-30 seconds run through the belts.
Hemp variety plays a key role in threshing efficiency. Make notes of any hemp lines that have unique characteristics (i.e., lack of shattered seeds before threshing).

Cloning

🧪Cloning🧪

Cuttings should be harvested from disease-free stock plants under non-stressed conditions. Collect turgid cuttings during optimal water conditions (non-wilted plants). Environmental conditions that increase propagation success are provided by an atmosphere that reduces water loss and maintains leaf turgidity, with optimal humidity between 75 and 90%, often achieved using humidity domes or “mini greenhouses”. Ample but not excessive light and clean, moist, and well aerated rooting media should also be provided (Casillas 2016). Ideal temperature should be around 25-27 °C (78-80 °F) with a root zone temperature of about 27-30 °C (80-85 °F). Once cuttings are taken, check for disease daily along with moisture content of media. If using humidity domes, monitor humidity levels and include fresh air daily.

Materials

Scissors/Pruning shears
Lab Gloves
IBA based rooting Hormone (we use Clonex™)
Beakers/Cups
Spray Bottle w/ water
Sterile medium (potting soil/Oasis Cubes/Water)
Humidity domes (if using potting soil + trays)
70% ethanol (alternative isopropyl alcohol)
Paper towels
Seedling flats
Disease-free Cannabis plants (ideally in the vegetative state)
Aeroponic apparatus
Mist timer
Heating mat

Common cloning tools and supplies

Methods (Aeroponic/Hydroponic)

Start with clean water (DI or tap) with no fertilizer
Cut lateral branch close to a node at a 45-degree angle
Gently scrape the stem around the cut to expose cambium layer
Remove any lower branches, keep approximately 3-5 nodes worth of leaves, leaving cutting looking like a “palm tree”
If taking many cuttings at a time, place cuttings in a beaker/cup of water until ready to dip into rooting hormone
Dip bottom 2 cm of cutting in rooting hormone, IBA (Clonex™)
Place cutting in aeroponic/hydroponic system
Let sit and watch for roots (7-14 days)
Use alcohol to disinfect tools before moving on to the next accession

Aeroponic cloning setup at PGRU

Methods (Potting soil/Oasis Cubes/Humidity Dome)

Cut lateral branch close to a node at a 45-degree angle
Gently scrape the stem around the cut to expose cambium layer
Remove any lower branches, keep approximately 3-5 nodes worth of leaves, leaving cutting looking like a “palm tree”
If taking many cuttings at once, place cuttings into beaker/cup filled with water until ready to dip into rooting hormone
When ready to root cuttings, emerge bottom 2cm of cutting in rooting hormone (Clonex™)
Place cutting with rooting hormone into seedling tray with potting soil or oasis cubes
Place humidity dome onto tray, mist water on inside of humidity dome
Let sit and watch for roots (7-14 days)
Use isopropyl alcohol to disinfect tools before moving on to the next accession

Typical humidity dome setup

Modified aeroponic rooting system protocol (Regas et al. 2021)

Generation of a mother plant for clonal propagation
1. Select healthy, female mother plant that exhibits desirable traits
2. Allow mother plant to reach the appropriate size (roughly 25 mature shoots) for clonal propagation
3. Allow mother plants to remain in the vegetative growth stage (light:dark = 18h:6h) to promote shoot growth for future propagation
Construction and preparation of aeroponic system
1. Begin by positioning the lid on top of the container. Drill desired number of holes into the lid while providing adequate space (~3cm) between each
2. Position water pump in the center of the container
3. Pour 7-8 L of distilled water into the container so that the pump nozzle remains roughly 2.3 cm above the waterline. NOTE: This ensures the submersible water pump can push water with enough force to spread across the container lid. Distilled water is recommended; however, regular tap water may also be used.
4. Situate the appropriate amount of Oasis Cubes or media of choice into each slot. Turn on the pump and allow it to run for 24 h on a set timer.
Selecting and excising appropriate shoots
1. Collect shoots near the apical meristem using a sterilized scalpel or scissor. Cuttings are ~10 cm in length, ideally with several nodes. NOTE: Cut the stem at a 45° angle. Cutting at a 45° angle increases the surface area of the basal portion of the cutting, allowing more space for root development.
2. Remove all foliage except foliage present on the top three nodes
3. Dip the newly excised cutting into the rooting solution containing indole-3-butyric acid (IBA) ~ 2-5 cm up from the base of the stem for ~5 s
4. Insert the cutting into the center of an Oasis cube positioned in the aeroponic system. NOTE: the cutting insertion depth is to remain ~1-2 cm from the bottom of the Oasis Cube
5. Mist the unrooted cuttings with the water every 100 seconds for 20 seconds
6. Grow the cuttings with 18-24 h of light per day with a photosynthetic photon flux density (PPFD) of 100 µmol/m2/s at 24-29°C and 40-60% relative humidity.
Aeroponic system maintenance and propagule health
1. Replenish the system with water at a pH between 5.0-6.0 every 2-5 days
2. Lightly mist the cuttings every 100 seconds for 20 seconds
3. Add 5 mL of each nutrient solution to the reservoir every 3-5 days
4. Add 15 mL of the algae and bacteria cleaning solution containing hypochlorous acid (0.028%) per 10 L of water every 5 days
Transplanting propagules
1. Select the cuttings with long, white, fibrous roots. NOTE: Avoid cuttings with brown, slimy, and short root systems as this is an indicator for the presence of root rot and will usually take longer to acclimate to the new growing medium and can bring unwanted diseases.
2. Place cutting into potting soil media, transplant propagules to 4 L nursery pot filled with a nutritious soil mix. NOTE: Watering immediately is recommended to prevent the roots from drying out.
Cleaning and storage of aeroponic system
1. When the system is no longer in use, wash with water and clean with 70% ethanol or another disinfectant (i.e Greenshield)
2. Remove the filter from the water pump and rinse with water to remove debris
3. Dry the system by wiping it down with paper towels or washcloth
4. Place pump inside the tub with the lid on and store until it is needed.

Tissue Culture

🧪Tissue Culture🧪

PGRU is extremely grateful to Faith Sparks for the development of this protocol. $ddH_{2}O$: deionized, distilled water.

Explant Collection, Preparation, and Induction:

Choose unlignified, young, vegetative, preferably apical nodal tissue as explants for in-vitro propagation.
Selectively cut the stem on an angle at a node to reduce water collection and to preserve structural integrity of the mother plant. Be sure to consider the direction of the apical node left on the mother plant to encourage growth in the direction most conducive to your growing environment.
Be sure to take cuttings that have at least three-six meristematic nodes and are 10 to 15 cm (4-6”) in length.
Always use clean and sharp shears.
Work from cleanest facilities to most infected to reduce pathogen spread.
Clean shears with ethanol if possible before moving on to the next plant.
Avoid taking cuttings from any visibly infested plants. Modify sterilization procedure to accommodate pest pressure.
Remove all fan leaves at least ½-1 inch from the stem leaving the length of the petiole (this will serve as a protective measure for the meristematic tissue against the destructive effect of the ethanol and bleach used in the sterilization process)
Place basal end of the cutting in clean water. Enclose the container if possible. Return nodal tissue to the laboratory as quickly as possible.
When transporting fresh cuttings, it is helpful to place the vessels in a cooler with ice to maintain the cuttings’ freshness as well as to halt bacterial growth at the cut sites prior to sterilization and in-vitro induction but this is not necessary for successful explant initiation.
Always ensure the cuttings’ basal ends are placed in vessels containing water for transport back to the laboratory. Plants will not survive the sterilization procedure if they have lost too much turgor pressure. Try to keep the humidity high in the vessel and avoid direct light or heat as these abiotic stressors may influence turgor pressure and ultimately make the cuttings more susceptible to unreconcilable damage.
It is best to sterilize and induct cuttings in-vitro within hours of sample collection, but it is possible to hold the fresh cuttings at 4 °C with ample relative humidity overnight if necessary. Do not sterilize cuttings prior to storage if they are to be held at 4 °C but do remove all foliage to reduce transpiration and to conserve space.
Do sterilize cuttings directly proceeding in-vitro induction after overnight storage.
After sterilization, further separate meristematic groupings with a sterile sharp scalpel so that each explant has about two meristematic buds per 2” of stem under a flow hood in a sterile environment with sterile tools.
Place explants in media in the pattern shown in figure III.
Replace vessel lid and place vessel in low light environment at 28 °C and 24-hour light.

Sterilization:

Spray down work surface with at least 70% ethanol and allow to dry.
If available, use a UV light to reduce the population of additional microbes. Some would say this is unnecessary, however, it is the technician’s preference. UV light will damage plant tissue so do not leave plant tissue exposed to UV light.
Sterilize all tools with flame or other heat source and use sterile glass plates for a cutting surface. The recommendation is to use 125 mm glass Pyrex dishes wrapped with aluminum foil and lined with sheets of 125mm Whatman paper. Autoclave for 20 minutes with a 20-minute dry cycle or comparable cycle to ensure sterility.
If desired or necessary, bring microscope into flow hood as cleanly as possible.
Submerge cuttings in 70% ethanol and seal the vessel under the flow hood. Apply gentle agitation for a duration of 2-2.5 minutes. Be sure to provide wiggle-room for the cuttings to be adequately sterilized on all surfaces.
Drain away the ethanol and rinse the cuttings once with sterile ddH2O.
Submerge the cuttings in 0.75-1.25% sodium hypochlorite (dependent on hemp variety and level of pathogen/pest present on the mother plant tissue) supplemented with 1-2 drops of tween per 100 mL of bleach solution as needed (ie. If there was a recent spray in the greenhouse or if there is substrate present on the plant tissue). Seal the vessel and apply gentle (<50rpm) agitation for ten minutes.
Immediately drain away the bleach/tween solution and rinse 5 or more times with sterile ddH2O until the smell of bleach is no longer detected and there is no more sodium hypochlorite left on the plant tissue.

Media Composition and MS Recipe Templates

Remove the vessel lid and pour clean and/or sterile H2O (approx. 50 mL) on the media.
Replace the covering ajar.
Change the water once or twice a day for three days and allow the vessel closure to be more and more open each day.
Pull plantlet out of the media and gently remove excess media on the roots.
Plant the rooted plantlet in LM-111 premoistened soil mix and water in with clean water.
Cover the tray with a tall dome and keep humidity high within the dome.
Place in a cool (but not less than 25 °C) spot of the greenhouse out of direct light for one week and maintain humidity and circulation consistently (avoid spikes and dips in RH and for photoperiod sensitive plants maintain vegetative growth with short nights and long days dependent on genotype).
Plantlets have survived omissions of the first two steps in this section, but acclimatization success rate is higher when a gradual humidity acclimatization is performed.

Best Practices

Always give newly inducted tissue a gradual introduction to high light environments. The reduction of shock is necessary after the sterilization procedure.
Never leave visibly damaged tissue on the cuttings after sterilization.
Use best judgment when transferring tissue early in-vitro and allow time and labor to be spent removing dead and dying tissue. If the tissue is left in the culture, it is believed ethylene may build up and cause the plants to senesce and/or flower in culture.
Always truncate cuttings to two-three nodes per explant.
Be sure your scalpels, forceps, and tool rests are all sterile and cooled to room temperature before allowing them to encounter sterile plant tissue, (hot tools cause unreconcilable damage to the delicate plant tissue and should not be used to handle the explants). * The author suggests using #10 blades and developing a cutting technique that leaves cleaner cut sites.
If there is any doubt the sterile explant has touched an unsterile surface discard explant immediately and re-sterilize the tool it is not worth contaminating a whole box or the rest of the sterile plant tissue.
Do not place more than five explants in each box if you can avoid it or the shoots will develop etiolated tissue that cannot be easily acclimatized back to the greenhouse, it is best to provide adequate space for shoots to receive a sufficient daily light integral. * Whenever possible, chose to use ventilated culture vessels. Providing sufficient gas exchange allows the tissue to develop more normally (ie. not hyperhydric) and saves time and resources throughout the process. Keep in mind ventilated lids make the cultures more susceptible to pathogen invasion.
Sow initial cultures onto Hemp TC media for one week-one months depending on the number of transfers that are necessary in early explant induction.
Long-night genotype cultures can be maintained on Hemp TC media indefinitely under a 24-hour photoperiod at approx. 50-90 $\mu$ mol m^{-2} s^{-1{} at 28 °C without flowering. Not all varieties of hemp are well suited to this environment, though and this may differ according to photosensitivity of the genotype.
Depending on contamination presence cultures may require additional sterilization procedures throughout the cultures in-vitro lifespan. It may be necessary to shorten the length of time between subcultures to reduce the population of the contaminant.

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