This manuscript reports several experiments that were conducted to develop a robust cryopreservation protocol for the long-term preservation of Cannabis sativa germplasm, a validation experiment to test the protocol across 13 genotypes, and a comparison of plant performance between cryopreserved material vs. non-cryopreserved plants. It should be noted that the experiments were conducted using the material that was available to the researchers at the time based on production schedules and other factors. As a result, some experiments were conducted with single genotypes while others were conducted with multiple genotypes, some the genotypes used for various steps were not always the same, and some steps were conducted in parallel rather than sequentially.
2.2. Cryopreservation Protocol
2.2.1. Conditioning
Explants were placed in preculture solution (PCS) dispensed in 62 × 95 mm baby food jars (hereafter referred to as “jars”) (Phytotech Labs Inc., Lenexa, KS, USA). PCS consisted of full-strength MS basal salts (Sigma-Aldrich Canada Co., Oakville, ON, CAN) and 0.5 M sucrose. PCS was filter-sterilized using a 0.20 µm polyethersulfone membrane sterilization unit (VWR™, Radnor, PA, USA). Explants were incubated for approximately 17 h under standard tissue culture room conditions while being agitated at 155 rpm on an orbital mini shaker (VWR™, Radnor, PA, USA).
Following the initial incubation period, explants were collected in a sterile, 40 µm nylon mesh cell strainer (VWR™, Radnor, PA, USA) while the PCS was allowed to flow through and be discarded. Using sterile forceps, explants were transferred back into their original jar or to a pre-sterile, 3 mL Neptune® polypropylene cryogenic vial (hereafter referred to as “vial”) (Neptune Scientific, San Diego, CA, USA). Loading solution (LS) was then added to the jar or vial at approximately 50- and 2-mL volumes, respectively The composition of LS was full-strength MS basal salts, 0.5 M sucrose, and 1.9 M glycerol (≥99.5% purity; Sigma-Aldrich Canada Co., Oakville, ON, CAN), filter-sterilized as previously described. Explants were incubated in LS for 20 min while being agitated at 155 rpm.
2.2.2. Vitrification
Nodal explants from position 2–4 were used to compare standard vitrification in pre-sterilized cryogenic vials to droplet vitrification. This initial comparison was conducted using plant vitrification solution 3 (PVS3), while later experiments were done using standard vitrification to compare PVS3 with plant vitrification solution 2 (PVS2) after it was identified to be more suitable.
2.2.3. Vitrification—Droplet
Following incubation in loading solution, explants were placed back into their original jar and submersed in approximately 50 mL of plant vitrification solution 3 (PVS3). Incubation in PVS3 occurred for 40 min with shaking at 155 rpm. While explants were incubating, autoclaved aluminum foil strips (hereafter referred to as ‘strips’) (approximately 0.5 × 2 cm) were aseptically prepared on 100 × 15 mm borosilicate glass petri dishes (VWR™, Radnor, PA, USA). Using a 5 mL polyethylene transfer pipette (VWR™, Radnor, PA, USA), single droplets of PVS3 were placed on the dull side of the strip (five drops per strip). Once incubation was complete, individual explants were placed into each of the droplets (see
Figure 2A). Strips loaded with PVS3 and explants were then plunged into LN at a slight downward angle. Strips were held in the LN for a few seconds to ensure that the entirety of the unit was sufficiently frozen and then remained in LN for at least 40 min.
Approximately 25 mL of US was dispensed into sterile jars and brought to 40 °C in a hot water bath (VWR™, Radnor, PA, USA). After at least 40 min in LN, the strips containing explants were transferred into 40 °C unloading solution (US) for 30 s. US contained full-strength MS basal salts, 0.8 M sucrose, and was filter-sterilized as described previously. The jar was swirled to release the explants from the strip, and the strip was removed immediately after the explants were freed. After 30 s, approximately 25 mL of room temperature US was added to the jar to bring the solution containing the explants closer to room temperature. Explants were incubated in US for 30 min on a rotary shaker at 155 rpm.
2.2.4. Vitrification—Conventional
For conventional vitrification using cryogenic vials, LS was removed and replaced with 2 mL of either plant vitrification solution 3 (PVS3) (used in earlier experiments, including droplet vs. conventional trial) or PVS2 (used in later experiments after PVS2 was found to perform better). The composition of PVS3 was full-strength MS basal salts, 50% sucrose, and 50% glycerol. PVS2 included full-strength MS basal salts, 0.4 M sucrose, 30% glycerol, 15% ethylene glycol (Fisher Scientific Company, Ottawa, ON, CAN), and 15% dimethyl sulfoxide (DMSO) (Phytotech Labs Inc., Lenexa, KS, USA). Both solutions were filter-sterilized as previously described. The vials were incubated at 155 rpm for 5 min before the solution was removed and replaced with 2 mL of fresh vitrification solution (VS). From here, vials were incubated while being shaken at 155 rpm for a predetermined amount of time (20–80 min). Following this incubation period, vs. was discarded and vials were replenished with 0.5 mL of fresh VS. Vials were immediately submerged in liquid nitrogen (LN; supplied by Linde Canada Inc., Mississauga, ON, CAN) and held there for at least 30 s with forceps to ensure adequate freezing of the explants. Vials were kept in LN for at least another 40 min (see
Figure 2B).
Approximately 50 mL of distilled water was dispensed into jars and brought to 40 °C in a hot water bath. After at least 40 min in the LN, vials were removed and immediately placed in jars containing the preheated water. Jars were continually swirled for 90 s to allow the explants to thaw. Subsequently, the vials were transferred to new jars containing approximately 50 mL of room temperature water. Jars were swirled over the duration of 60 s before the vials were removed. vs. was discarded from the vials and replaced with 1 mL of US. Vials were incubated for 30 min on a rotary shaker (155 RPM).
2.2.5. Recovery Preparation
After the unloading step, US was removed by either passing through a cell strainer into a waste bottle (the strainer collecting explants from jars) or by use of a pipette (leaving only explants in the vials). Using sterile forceps, explants were transferred to autoclaved, ashless filter paper to blot dry. Subsequently, explants were plated onto recovery media, right-side up (or on their side if orientation could not be discerned). Please see
Figure 2C for a visual representation of samples plated on recovery media.
2.2.6. Recovery and Data Assessment
Samples were allowed a recovery period of 30 days in culture. Thereafter, growth was assessed and categorized in terms of survival and regeneration. ‘Survival’ was classified as explants that remained green but may not have shown visible growth. ‘Regrowth’ was demonstrated by leaf development on explants (see
Figure 2D). Samples were analyzed using a stereo microscope (Leica Microsystems, Wetzlar, Germany). For all cryopreservation experiments, three treatment groups (‘control’, ‘no LN2’, ‘LN2’) were compared. ‘Control’ samples were excised from the donor plant and plated directly onto recovery media to determine the baseline survival rate of the explants. ‘No LN2’ samples were exposed to the cryopreservation protocol but were not frozen in LN to assess the effect of pretreatments on explant health. ‘LN2’ samples were processed through the entire cryopreservation protocol including the freezing process. At least 10 explants were situated on each plate, and each cryopreservation and recovery media treatment were replicated at least two times. Contaminated plates were removed from the data when performing statistical analysis.
2.2.7. Growth Conditions for Recovering Samples
Samples were incubated in the dark in a culture room (24 ± 2 °C) for a predetermined amount of time before either gradual or rapid exposure to ambient light conditions (16/8-h light/dark photoperiod) under cool white fluorescent or LED lighting as previously described.
2.2.8. Experiment—Recovery Media
A variety of recovery media were investigated using cryopreserved Cultivar 1 samples. The media tested included SMM, half strength SMM (HalfSMM), which was the same as SMM except made with half-strength MS basal salts with vitamins, HalfSMM with supplemental GA3 (HalfSMM + GA3) which was composed of HalfSMM with the addition of 1 μM Alfa Aesar™ gibberellic acid (GA3; Fisher Scientific Company, Ottawa, ON, CAN), MS basal medium (MSbasal) composed of full-strength MS basal salts with vitamins, 30 g L−1 sucrose, 0.3 g L−1 activated charcoal, 8 g L−1 agar, and MSbasal with GA3 (MSbasal + GA3) composed of MSbasal supplemented with 1 μM GA3.
GA3 was prepared as a 100 mM working stock solution by dissolving the powder in 99% ethanol before diluting it with distilled water. All media were pH adjusted to pH 5.7 ± 0.2 before the addition of agar and autoclaved before being aliquoted into vessels as described previously. Approximately 25 mL of autoclaved media was dispensed into pre-sterile, 100 × 15 mm plastic petri dishes (VWR™, Radnor, PA, USA) under aseptic conditions.
Ten explants were used per treatment and each treatment was replicated thrice. Sample response to recovery media were assessed using only the droplet vitrification protocol for SMM, HalfSMM, and HalfSMM + GA3, while both strip and vial protocols were performed for the recovery of Cultivar 1 explants on SMM, MSbasal, and MSbasal + GA3 media. All samples were cryopreserved using PVS3 with an exposure time of 40 min. The comparison of multiple genotypes responding to the cryopreservation protocol was performed only using MSbasal for the recovery media.
2.2.9. Experiment—Cold Incubation of Donor Plants
The effect of cold incubation of donor plants before use for cryopreservation was investigated using Cultivar 2. Briefly, half the cultures were placed into an incubator (Norlake® Tissue Culture Chamber Model; Standex International Corporation, Salem, NH, USA) programmed to 10 ± 1 °C (16/8-h day/night photoperiod, approximately 42 µmol/m2/s) after 4 weeks of growth. These cultures were maintained for 7 days at this temperature while the other half remained at 24 ± 2 °C. Both sets of cultures were used for cryopreservation at week 5.
Ten explants were used per treatment and each treatment was replicated thrice. Samples were cryopreserved using the conventional cryopreservation protocol and a 40-min exposure to PVS3. Samples were plated onto MSbasal recovery medium for recovery. Samples recovered under one of two light regimes: darkness for 7 days, gradual build-up to ambient light intensity for 7 days, then ambient light intensity for 16 days; darkness for 7 days, then ambient light intensity for 23 days. Light was provided by programmable LEDs. Sample survival and recovery were assessed 30 days after plating.
2.2.10. Experiment—Extension of Incubation in Darkness
The response of cryopreserved Cultivar 1 samples to increased incubation in darkness during the recovery period was investigated. Ten explants were used per treatment and each treatment was replicated thrice. Samples were cryopreserved using the conventional cryopreservation protocol and a 40-min exposure to PVS3. Samples were plated onto MSbasal recovery medium for recovery.
Light treatments included the following:
- (1)
Darkness: 5 days, gradual light: 5 days, ambient light: 20 days
- (2)
Darkness: 10 days, gradual light: 5 days, ambient light: 15 days
- (3)
Darkness: 15 days, gradual light: 5 days, ambient light: 10 days
- (4)
Darkness: 20 days, gradual light: 5 days, ambient light: 5 days
- (5)
Darkness: 7 days, gradual light: 7 days, ambient light: 16 days
- (6)
Darkness: 14 days, gradual light: 7 days, ambient light: 9 days
- (7)
Darkness: 14 days, gradual light: 0 days, ambient light: 16 days
- (8)
Darkness: 21 days, gradual light: 7 days, ambient light: 2 days
During the period of gradual light exposure for treatments 1–4, five sheets of white printer paper were placed on top of the cultures. One sheet of paper was removed each day for the five-day period, which was succeeded by the samples being subjected to normal culture room light intensity. For samples treated with light treatments 5–8, the light source was programmed to slowly increase in magnitude for the duration of the gradual light intensity stage until ambient levels were achieved. Light was provided by cool white fluorescent bulbs (treatments 1–4) or programmable LEDs (treatments 5–8).
2.2.11. Experiment—Comparison of PVS3 and PVS2
The recovery of Cultivar 2 samples after cryopreservation using either PVS3 or PVS2 in cryogenic vials was investigated. Ten explants were used per treatment and each treatment was replicated thrice. The preliminary exposure time to PVS3 (40 min) was compared to PVS2 exposure times of 20, 30, 40, 60, and 80 min. Samples were plated onto MSbasal recovery medium and allowed to recover under darkness for 5 days, followed by low light conditions (5 days) then ambient light (20 days). Sample survival and recovery were assessed 30 days after plating.
2.2.12. Growth Conditions for Donor Plants and Recovered Cultures
Samples that recovered from cryopreservation (demonstrating at least new leaves) were subcultured on either SMM or SMM supplemented with 2 µM GA
3 (SMM + GA
3) and were allowed to grow for at least four weeks. Cultures grew under a 16/8-h light/dark photoperiod at 24 ± 2 °C. Light was provided by either cool white fluorescent bulbs or LEDs (see
Figure 3A) and emitted a photon flux of 42–52 µmol/m
2/s.
2.2.13. Experiment—Cultivar Response to Cryopreservation Protocol
The optimized cryopreservation protocol was used to assess the survival and regrowth of 13 commercial cannabis genotypes. Briefly, samples were vitrified using conventional vitrification using an exposure period of 60 min in PVS2. Samples were recovered on MSbasal medium in the dark for five days followed by 25 days at 42 µmol/m2/s. Ten explants were used per cultivar and each cultivar was replicated thrice. Survival and recovery were assessed at 30 days post-cryopreservation.
2.2.14. Evaluation of Cryopreserved Plants
A subsample of control (
n = 4), no LN2 (
n = 2), and LN2 (
n = 2) Cultivar 1 cultures from early trials (droplet vitrification method, 40-min PVS3 exposure, gradual exposure to light supplied by cool-white fluorescent bulbs) were selected for hardening off and ex vitro growth (see
Figure 3B,C). These samples were used for subsequent phenotyping and measurement of specific chemical compounds commonly detected in cannabis inflorescence (hereafter referred to as “bud”) and trim material. Cultures that were selected for ex vitro growth displayed visually normal morphology (explants developing elongated shoots) and had developed adventitious roots while in culture without a specific rooting treatment.
The plants were transferred out of culture on 26 June 2018. Approximately 15 mL of autoclaved tap water was poured into the vessel of each plant being transplanted and the media was allowed to soften. The plants were subsequently removed from the vessels and the roots were separated from the medium under running tap water. A small amount of tap water was added to each vessel along with their respective sample to ensure that the roots would not dry out.
Individual plants were planted in 500 mL pots, which were filled with moist, autoclaved, soilless potting mix (Pro-Mix HP
® Mycorrhizae, Pro-Mix, Rivière-du-Loup, QC, CAN). Briefly, the roots of the plants were placed into the small hole created in the center of each pot and covered with the potting mix (see
Figure 3D). Once the plants had been transferred, they were watered with approximately 80 mL of diluted half-strength vegetative fertilizer. The pH and EC of the veg feed were approximately 6.5 and 1.0, respectively. Plants were watered with this for one week after transplanting. For the first 96 h, the plants were given 80–100 mL of feed daily. Following that, the plants were watered as needed. Beginning in the second week of hardening, the plants were watered with veg 1 solution (EC 2.0).
The potted plants were placed into a clone tray (27.8 × 54.5 × 6.2 cm; T.O. Plastics®, Clearwater, MN, USA), transferred to a clone propagation cart located in an environmentally controlled grow room, and covered with a humidity dome. The vents on the humidity dome were opened after 48 h, and the dome was removed 96 h after transplanting. For 7 days, the light levels were set to half of the maximum intensity (approximately 150 µmol/m2/s), supplied by T5 fluorescent bulbs and programmed to an 18/6-h day/night photoperiod. After the initial week, the light intensity was increased to approximately 250 µmol/m2/s.
2.2.15. Vegetative Growth
After new growth was observed and roots were seen protruding from the sides of the pots, the plants were transferred to 3.79 l pots filled with growing medium and relocated to a grow room for vegetative growth (week of 9 July 2018). The plants were allowed to grow for approximately 2 months under an 18/6-h day/night photoperiod and light supplied by high pressure sodium (HPS) and metal halide (MH) lights. Dead and yellowing leaves and lateral stems in close proximity to the base of the plant were removed as needed to ensure adequate air flow and reduce the risk of disease. Plants were watered with pH- and EC-adjusted nutrient solution as needed via drip irrigation.
2.2.16. Flower Induction
On 12 September 2018, plants were subjected to a 12/12-h light/dark photoperiod for floral induction. Light was supplied by HPS and MH ballasts. Plants were watered with pH- and EC-adjusted nutrient solution as needed via drip irrigation. The plants were subjected to a nutrient flush with water one week before harvest (see
Figure 3E).
2.2.17. Bud and Trim Harvest
On 7 November 2018, whole plants were cut at the base of the stem (just above the medium) and whole plant weights were taken immediately. Lateral branches were cut from the main stem and both trim and bud tissues were removed from the branches. The combined weights of the main stem and lateral branches were subsequently taken.
Fresh bud was hand-trimmed to remove any additional fan leaves and stem; these tissues were added to the trim material. Weights of freshly harvested plants, bud, and trim were recorded; fresh stem weights were calculated by subtracted fresh bud and trim weights from the total plant weight. Fresh bud and trim harvested from individual plants were then transferred to a drying room (16–21 °C, 35–65% relative humidity), spread over stainless steel screens (Bundy Baking Solutions, Urbana, OH, USA) placed on drying racks (Metro® 2660 Dry Unit, Metro Shelving, Curtis Bay, MT, USA) and allowed to dry for at least 7 days.
2.2.18. Moisture Analysis
Before harvested plant material could be packaged, moisture content was measured using a Mettler Toledo® HE73 Moisture Analyzer (Mettler Toledo, Mississauga, ON, CAN) with a run temperature set to 95 °C. A subsample of bud weighing 3–5 g (exact weight recorded) was milled using a hand grinder onto an aluminum pan and loaded into the heating module. Sample moisture content was expressed as a percentage once drying had commenced. The harvested product could be packaged and used for further analysis once moisture content had reached <9%. Product was packaged in a plastic packaging pouch that was either heat (Uline® Tabletop Poly Bag Sealer, Uline Canada, Milton, ON, CAN) or vacuum (Henkelman Vacuum Systems® Boxer 42 XL, Henkelman BV, CJ ‘s-Hertogenbosch, Netherlands) sealed and subsequently stored at room temperature.
Dry weights for the bud and trim from each plant were recorded immediately before packaging and bud and trim moisture content (%) and bud yield per plant (g/g dry weight) were calculated.
2.2.19. Cannabinoid and Terpene Detection and Quantification
Approximately 10 g of bud and trim sample (actual weight recorded) was submitted for measurement of cannabinoids and terpenes for each individual plant.
For cannabinoid analysis, samples were homogenized using a mortar and pestle. 0.5000 g (+5% tolerance) of milled sample was weighed and transferred to a 10 mL test tube. Extraction solution (10 mL) was added to each sample. Each sample was then vortexed for 30 s, sonicated for 30 min, then vortexed a second time. The supernatant from each sample was transferred to a dilution vial (A) using a 3- or 10-mL glass or plastic 0.2 µm filter syringe after centrifugation for 3 min at 1000× g. The first couple milliliters of filtrate was discarded and not used for analysis in case cannabinoids were bound to the membrane. 50 µL of sample from dilution vial A was transferred to a second dilution vial (B). 900 µL of dilution solution was added to dilution vial B along with the sample. Cannabinoids were measured using an Agilent Technologies High Performance Liquid Chromatography (HPLC)—1200 Infinity system with a diode array detector (DAD) (Agilent Technologies Inc., Santa Clara, CA, USA).
For terpene analysis, approximately 5 g of sample was homogenized in a mortar and pestle and transferred to a 15 mL plastic centrifuge tube. Approximately 500 mg (+2% tolerance) of homogenized sample was loaded into the headspace vial directly. Terpenes were measured using an Agilent 7820A/7890B gas chromatograph (GC) system with flame ionization detection (FID) (Agilent Technologies Inc., Santa Clara, CA, USA). Data obtained from the samples was analyzed by Chemstation® software (Open LAB CDS Chemstation Edition Rev. A.02.02(1.3), ChemStation International Inc., Dayton, OH, USA).
A total of nine cannabinoids and 23 terpenes were investigated for this study. Peaks on the chromatographs were identified by internal (in-house partially decarboxylated bud material) and external cannabinoid (Cerilliant Corporation, Round, Texas, USA) and external terpene (Sigma-Aldrich Canada Co., Oakville, ON, CAN; Restek Corporation, Bellefonte, PA, USA) standards. Final values are provided as %w/w of the original dried material (see
Supplemental Figures S1 and S2 for HPLC-DAD and GC-FID chromatographs, respectively).