Journal:Cadmium bioconcentration and translocation potential in day-neutral and photoperiod-sensitive hemp grown hydroponically for the medicinal market
| Full article title | Cadmium bioconcentration and translocation potential in day-neutral and photoperiod-sensitive hemp grown hydroponically for the medicinal market |
|---|---|
| Journal | Water |
| Author(s) | Marebesi, Amando O.; Lessl, Jason T.; Coolong, Timothy W. |
| Author affiliation(s) | University of Georgia |
| Primary contact | Email: aom at uga dot edu |
| Year published | 2023 |
| Volume and issue | 15(12) |
| Article # | 2176 |
| DOI | 10.3390/w15122176 |
| ISSN | 2073-4441 |
| Distribution license | Creative Commons Attribution 4.0 International |
| Website | https://www.mdpi.com/2073-4441/15/12/2176 |
| Download | https://www.mdpi.com/2073-4441/15/12/2176/pdf?version=1686282553 (PDF) |
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This article should be considered a work in progress and incomplete. Consider this article incomplete until this notice is removed. |
Abstract
Heavy metal contamination of agricultural soils is potentially concerning when growing crops for human consumption. Industrial hemp (Cannabis sativa L.) has been reported to tolerate the presence of heavy metals such as cadmium (Cd) in the soil. Therefore, the objectives of this study were to evaluate Cd uptake and translocation in two day-length-sensitive (DLS) and two day-neutral (DN) hemp varieties grown for the medicinal market and to determine the impact of Cd exposure on cannabinoid concentrations in flowers. A hydroponic experiment was conducted by exposing plants to 0 mg·L−1 Cd and 2.5 mg·L−1 Cd in the nutrient solution. Cadmium concentrations ranged from 16.1 to 2274.2 mg·kg−1 in roots, though all four varieties accumulated significant concentrations of Cd in aboveground tissues, with translocation factors ranging from 6.5 to 193. Whole-plant bioconcentration factors ranged from 20 to 1051 mg·kg−1. Cannabinoid concentrations were negatively impacted by Cd exposure in DN varieties but were unaffected in DLS varieties. Biomass was reduced by Cd exposure demonstrating that these varieties might not be suitable for growth on contaminated soil or for phytoremediation. There is potential for Cd accumulation in flowers, showing the need for heavy metal testing of C. sativa consumer products.
Keywords: auto-flower, bioconcentration factor, cannabinoid, heavy metal, hemp, metal stress, translocation factor
Introduction
For centuries, humans have cultivated hemp (Cannabis sativa L.) for its fiber, seed, therapeutic, and psychoactive properties. During the domestication process, wild Cannabis populations have been subject to selection, giving rise to the multiple varieties that exist today. [1] The term "industrial hemp" is commonly used to refer to C. sativa plants [2] with total tetrahydrocannabinol (THC) concentrations below 0.3%. Plants with total THC concentrations above 0.3% are classified as marijuana and subject to federal prohibition in the United States (U.S.) [3] Current industrial hemp breeding efforts target fiber, seed, or cannabinoid production, based on the end user. Hemp varieties with high cannabidiol (CBD) concentrations are often selected for medicinal and therapeutic uses. There is also a focus on developing varieties that are day-neutral (DN) or minimally sensitive to photoperiod in order to expand production opportunities. [4]
Hemp is generally considered a qualitative short-day plant that flowers in response to decreasing photoperiods. Hemp selections that flower in response to photoperiod are known as day-length-sensitive (DLS). After emergence, hemp undergoes a photoperiod-dependent vegetative phase maintained by exposure to approximately 14–18 hours or more of light daily. [5,6] When hemp is planted during periods of short days (<13 hours of light), it may flower prematurely. Premature flowering, prior to complete vegetative development, can result in yield reductions. [7,8] In contrast, some hemp varieties exhibit DN flowering tendencies known colloquially as “auto-flower” hemp. These DN varieties are relatively insensitive to photoperiod for flower induction. The DN trait is speculated to arise from Cannabis ruderalis (C. sativa ssp. ruderalis) and may have originated from hemp located in high latitudes where photoperiods can be long and growing seasons are typically short or regions with relatively short and constant photoperiods. [5,9,10] Advantages of DN hemp varieties include the ability to flower in regions that have little variation in photoperiod throughout the year (tropics) or during times of the year when photoperiods may be inadequate to grow DLS varieties. However, many DN types of hemp have been reported to be particularly sensitive to environmental stressors such as high temperatures and may have lower yields than comparable DLS varieties. [11]
In addition to uses for fiber, seed, and medicinal purposes, hemp has also been proposed as a candidate for use in phytoremediation, which utilizes plants to remove contaminants, such as heavy metals or other chemicals from soils. [12,13] Accumulator plant species can uptake heavy metals from soils, even at low external concentrations, and concentrate them in plant tissues. [14] By growing accumulator plants in contaminated soil, it is possible to realize in situ decontamination, an economically viable approach that preserves physicochemical soil characteristics, while removing contaminants. [15] The morphophysiological characteristics of hemp, such as high biomass production, deep roots, and short life cycle, make it a potential candidate for phytoremediation. [16,17,18,19,20]
Heavy metal contamination of agricultural soils is a concern when growing crops for food or medicinal purposes, due to potential harm to human and animal health. [21,22,23] Cadmium (Cd) contamination in the environment has been linked to anthropogenic activities, such as mining and smelting. Further, Cd can be introduced to soils via contaminated manure, sewage sludge, and phosphate fertilizers. [24] Cadmium is known to cause health issues when ingested in amounts greater than the provisional tolerable monthly intake (PTMI) of 25 μg·kg−1 of body weight. [25] In previous studies utilizing naturally and artificially contaminated soil and substrate containing from 0 to up to 200 mg·kg−1 Cd, hemp varieties grown for fiber production accumulated Cd in aboveground tissues at levels that could be harmful to human health. [26,27,28,29] For instance, the hemp fiber variety Silistrinski grown in naturally contaminated soil containing 12.2 mg·kg−1 Cd accumulated 1.22 mg·kg−1 Cd in its flowers. [27]
There are multiple indicators that can be used to determine the accumulation potential of a plant species. Bioconcentration factor (BCF) is the ratio between the metal concentration in plant tissues and the initial metal concentration in the soil or growing solution. [20,30,31,32,33] This indicator has also been used interchangeably with terms such as accumulation factor (AF) [28], biological absorption coefficient (BAC) or index of bioaccumulation (IBA). [23] A separate indicator of accumulation potential is the translocation factor (TF), which is the ratio between the metal concentration in the above ground biomass and the metal concentration in the roots. [31,32] Additionally, plant growth parameters can be assessed to determine the tolerance index (TI), calculated as the ratio between growth in contaminated and non-contaminated soils. [20,30] There is significant variability in BCF among plant species and chemical elements. It has been proposed that plants with BCF >100 mg·kg−1 Cd on a dry weight (DW) basis in its leaves could be referred to as hyperaccumulators. [14] Conversely, Chaney and Baklanov [12] suggested that true hyperaccumulators are able to accumulate higher concentrations of metals in leaves than in roots (TF > 1).
Few studies have evaluated heavy metal accumulation in hemp flowers, with most research utilizing fiber hemp varieties to determine heavy metal uptake for phytoremediation purposes. [16,17,19,34] Due to the harmful effects of Cd and other heavy metals on human health, the U.S. hemp industry has attempted to implement standards regarding maximum allowable levels of metals in C. sativa consumer products, which vary by state. [35] As hemp flowers are increasingly grown for the medicinal market, determining Cd distribution among plant organs, as well as bioconcentration and root-to-shoot translocation factors, are of importance. We hypothesize that there are distinctions in Cd accumulation and distribution among plant tissues in hemp varieties with different growth and flowering habits. Therefore, the objectives of this study were to evaluate nutrient partitioning and Cd uptake, translocation, and accumulation in DLS and DN hemp varieties, and to determine the impact of Cd exposure to cannabinoids in plant flowers.
Materials and methods
Experimental settings
The experiment was conducted in a greenhouse in Watkinsville, GA, USA (lat. 33°5′ N, long. 83°3′ W) from January 2022 to April 2022. Feminized seed from two DLS hemp varieties, T1 and Von (Sunbelt Hemp Source, Moultrie, GA, USA), and two DN varieties Apricot Auto (Blue Forest Farms, New York, NY, USA) and Auto CBD Alpha Explorer (Alpha Explorer) (Phylos Bioscience, Portland, OR, USA) were sown into engineered foam cubes (3.33 cm L × 2.54 cm W × 3.81 cm D; Oasis Grower Solutions, Kent, OH, USA) for germination. Foam cubes were placed in plastic trays over a germination mat set at 24 °C exposed to a mist irrigation system, which applied water twice daily for one minute each. Supplemental lighting (approximately 100 µmol·m−2·s−1) was used during germination. Seedlings were maintained under these conditions for four weeks, after which they were placed into plastic netted containers (4.7 cm W × 5.1 cm D) and transferred to 37.9 L plastic containers (Rubbermaid Inc. Wooster, OH, USA) filled with 28 L of well water. The well water was analyzed for nutrient concentrations periodically throughout the experiment (Table 1). Four seedlings per replicate were placed equidistantly (24.3 cm apart) in holes drilled in the container lid. Welded wire mesh frames were attached to each lid to support plants. A 15.2 cm aquarium air stone attached to an air pump (Active Aqua; Hydrofarm, Petaluma, CA, USA) was placed inside the container to aerate the nutrient solution throughout the experiment. Container volume was maintained by adding well water every two to three days. At transplant, a nutrient solution was added to the plastic containers using a half-strength Hoagland’s solution [36] (Table 1).
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Plants were grown for three weeks in the base nutrient solution after which nutrient solutions were replaced completely and Cd treatments were added using CdSO4·8H2O, to achieve 0 (control) and 2.5 mg·L−1 Cd. Cadmium concentrations were chosen based on the results of previous studies [34,35], which evaluated hemp exposure to Cd in hydroponic systems. The experimental treatments were arranged in a randomized complete block design, with four hemp varieties exposed to two levels of Cd with four replicates each. Nutrient solutions were maintained to a constant volume by adding water every two to three days, and nutrients were replaced every three weeks for the remainder of the experiment. The electrical conductivity (EC) and pH of the solutions were measured weekly. Solution pH was adjusted to 5.5 when necessary (pH down; General Hydroponics, Santa Rosa, CA, USA). Supplemental light (approximately 100 µmol·m−2·s−1) was used to provide 18/6 light/dark hours for four weeks of vegetative growth after transplanting seedlings into containers. Supplemental lights were turned off to allow for flower induction in the DLS varieties for the remaining seven weeks of production (average day length 12 hours and 38 minutes). The DN varieties (Apricot Auto and Auto CBD Alpha Explorer) exhibited visually detectable flower development one week prior to the induction of flowering in the DLS varieties (Von and T1). Therefore, the DN varieties were harvested one week prior to the DLS varieties to ensure that plants flowered for the same length of time.
Nutrient solutions were sampled at the beginning and end of each three-week cycle using 20 mL scintillation vials (HDPE; Thermo-Fisher Scientific™, Waltham, MS, USA), and stored at −4 °C until the analysis of mineral nutrient concentrations (Table S1). Temperature and relative humidity (RH) of the greenhouse were monitored at canopy height hourly (VP4; Meter Group Inc., Pullman WA, USA) and averaged 19.1 ± 3.1 °C and 74 ± 0.1% RH for the experiment. Photosynthetic active radiation was also monitored hourly throughout the experiment (QSO-S; Meter Group Inc.) and plants were exposed to an average daily light integral (DLI) average of 21.6 ± 9.0 mol·m−2·d−1.
Mineral analysis
Samples of fresh root, stem, leaf, and flower tissues were collected for Cd analysis at harvest. Composite samples (50 g fresh material) were taken from each of the four plants in a replicate (container). Roots were triple-washed with deionized water after removal. Ten of the youngest fully expanded leaves were collected from the top one-third of each plant (main stem and lateral branches) and rinsed with deionized water. Stem samples were collected from the bottom two-thirds of the main stem. Flower material was sampled from the top of the main stem and the top one-third of plants. Samples were placed in a forced air oven set at 55 °C for 72 hours until a constant weight was achieved. Dried plant material was then ground in a Wiley mill (Thomas Scientific, Swedesboro, NJ, USA) and passed through a 20-mesh screen. Samples were digested using EPA Method 3052. [37] In brief, 0.5 gram samples were placed in fluorocarbon polymer microwave vessels, 10 mL of concentrated nitric acid were added to each vessel which was then sealed. The microwave digester (Mars 6 Microwave; CEM Corp., Matthews, NC, USA) was heated to 200 °C for 30 minutes and digested (solutions) were then transferred quantitatively into volumetric flasks and brought to 100 mL volume with deionized water prior to analysis.
Samples of the hydroponic solutions were filtered using a 0.45 µM PTFE membrane (Thermo-Fisher Scientific™ Choice™ Polypropylene Syringe Filters) and acidified using 2% (v/v) high purity nitric acid (HNO3) (Certified ACS Plus, Fisher Scientific, Pittsburgh, PA, USA) prior to analysis. Hydroponic solutions and plant tissues were analyzed for multiple elements—phosphorous (P), potassium (K), sulfur (S), calcium (Ca), magnesium (Mg), iron (Fe), manganese (Mn), aluminum (Al), boron (B), copper (Cu), zinc (Zn), nickel (Ni), and cadmium (Cd)—following EPA Method 200.8 [38] by inductively coupled plasma–optical emission spectroscopy (Spectro Arcos FHS16; Spectro Ametek USA, Wilmington, MA, USA). The instrument parameter settings and wavelengths used are displayed in the supplementary material (Tables S3 and S4, and Figure S1). The instrument reporting limit for Cd was <0.005 mg·L−1. Results were expressed as mg·L−1. Calibration standards utilized in this analysis were from a certified source (Inorganic Ventures, Christiansburg, VA, lot number: N2-MEB667614). Independent laboratory performance checks were also run with acceptable deviations for recoveries set at 100 ± 5.0%.
The BCF was calculated by dividing the Cd concentration in plant tissues by the initial Cd concentration in the nutrient solution. [20,30,31,32,33] The TF (%) was calculated by dividing the sum of Cd concentration in leaves, flowers, and stems by the Cd concentration in roots, and multiplying by 100. [31,32] Cadmium uptake rates (µmol·plant−1·d−1) were calculated by the following equation (adapted from Ali et al.[30]): ((([Cdinitial − Cdfinal])/number of plants)/treatment days)/root biomass)
Plant growth and biomass yield
Plant height was determined by measuring the distance from the base of the stem to the tip of the apically dominant flower at harvest [11].
To quantify leaf, flower, stem, and root biomass, four whole plants per replicate were air-dried at ambient temperatures inside the greenhouse for two weeks and then separated into roots, stems, and leaf and flower biomass. Dry leaf and flower materials were manually pulled from plants following industry standards used for hemp biomass intended in cannabinoid extraction. Subsamples were taken from the air-dried materials and further dried in a forced air oven set at 55 °C for 48 hours until a constant weight was achieved. The dry weights of the whole plant samples were then normalized based on subsample moisture content. [11]
Cannabinoid analysis
Approximately 25 g of fresh flower tissue sampled from inflorescences located on the top one-third of the plants were sampled during weeks six (DN varieties) and seven (DLS varieties) of flowering (49 and 56 days after treatment [DAT], respectively) and dried separately from other samples as follows. Flower material was placed on a perforated aluminum baking sheet and dried to approximately 15% moisture content in a walk-in cooler with a temperature set point of 13 °C and 55% relative humidity for 14 days. The appropriate relative humidity was maintained using a dehumidifier. The dried material was hand trimmed to remove leaves, sealed in a metalized resealable food bag (Uline, Braselton, GA, USA) and stored at −4 °C for cannabinoid analysis. The acidic and neutral forms of the cannabinoids, THC and CBD, were determined in dried flower material by a commercial laboratory using high-performance liquid chromatography and a diode array detector set to 230 nm (SJ Labs and Analytics, Macon, GA, USA). The limit of detection for THC and CBD was 0.02%. Total cannabinoid concentrations were calculated by the following formula: total cannabinoid = neutral + (acidic form × 0.877). The percentage of dry matter for all samples was recorded and the results were reported on a dry weight basis.
Statistical analysis
Statistical analysis was conducted using JMP Pro 15 (SAS, Cary, NC, USA). Data were subjected to a one-way ANOVA procedure with Student’s t-test (p < 0.05) or Tukey’s Honest Significant Difference test (p < 0.05) conducted for mean separation when appropriate. Tissue Cd and cannabinoid concentrations were log-transformed to ensure equal variance prior to statistical analysis. Non-transformed data are presented.
Results and discussion
Plant height and biomass yield
References
Notes
This presentation is faithful to the original, with only a few minor changes to presentation. Some grammar and punctuation was cleaned up to improve readability. In some cases important information was missing from the references, and that information was added.
