CBD oil by-product (Hemp flakes): Evaluation for nutritional composition, heavy metals and functionality as a food ingredient
Department of Food and Animal Sciences, Alabama Agricultural and Mechanical University, Normal, AL, USA
Winfred Thomas Agricultural Research Station, Hazel Green, AL, USA
Department of Biological and Environmental Sciences, Alabama Agricultural and Mechanical University, Normal, AL, USA
∗Corresponding author. josh.herring@aamu.eduAbstract
Background
The recent interest among consumers in industrial hemp due to health and wellness benefits has led to several products from industrial hemp, including cannabidiol (CBD) oil. CBD oil extraction from hemp buds and flowers generates by-product biomass (hemp flakes), often posing disposal challenges and with little or no applications. We hypothesized that hemp flakes possess residual compounds with nutritional and health value that could be used to improve utilization.
Methods
Locally sourced hemp flakes were compared to three commercial hemp protein products. The nutritional composition (proximate analysis), heavy metals (Al, Cu, As, Pb, Co, Cd), and functional composition (phenolic and antioxidant properties–total phenolic compounds (TPC), total flavonoid compounds (TFC), ferric reducing antioxidant potential (FRAP), 1,1–diphenyl-1-picrylhydrazyl (DPPH), Trolox equivalent antioxidant capacity (TEAC)), (CBD, cannabiodiolic acid–CBDA, cannabichromene–CBC, cannabigerol–CBG, and cannabinol–CBN) contents were determined and compared.
Findings
Hemp flakes had a similar nutritional composition to commercial hemp protein products, with heavy metal levels within FDA allowed limits. The by-product had significantly higher CBDA levels than commercial products. Overall, hemp flakes had comparable nutrient composition and antioxidant capabilities. Based on the protein composition of hemp flakes (31.62 %) versus the highest commercial product (43 %), hemp flakes are an acceptable functional food ingredient.
Highlights
- •The nutritional profile of hemp flakes compared with commercial hemp protein products.
- •Heavy metal levels of hemp by-product were at FDA permissible levels.
- •Hemp by-product showed significantly high CBDA content compared to commercial hemp products.
- •The hemp by-product could be utilized as a functional food ingredient.
1Introduction
Industrial hemp (Cannabis sativa L.) is a versatile plant now cultivated globally for industrial, medicinal, and food purposes [1]. Unlike its counterpart, marijuana, industrial hemp contains less than 0.3 % of the psychoactive compound tetrahydrocannabinol (THC). Cannabidiol (CBD), a non-psychoactive compound, has shown therapeutic potential for treating conditions like body pain, anxiety, and insomnia [2]. Other reported benefits of industrial hemp include anti-inflammatory, antipsychotic, and anticonvulsant properties [3,4]. Its mechanisms of action are linked to its ability to mimic endocannabinoids that regulate the nervous system. These potentials have fueled consumer demand for CBD-infused products, prompting manufacturers to incorporate them into various available food and beverage items [5].
Hemp was not legally recognized in the United States until the 2018 Farm Bill [3], and it still has use limitations by state. Around the same time, regulatory bodies in the US and UK approved CBD-based medications for treating epilepsy and multiple sclerosis symptoms.
CBD is not allowed in foods on a federal basis [6,7]; however, some states have legalized recreational (including food applications) and medicinal use, while others approved it only for therapeutic use. CBD is extracted from industrial hemp through cold pressing or chemical extraction. The by-product of CBD extraction are hemp flakes. Hemp flakes contain residual CBD, antioxidant compounds, and nutritional value. The cannabionoids and antioxidiant compounds can account for functional properties. A functional ingredient is one that provides benefits to the consumer beyond nutrition and is capable of mitigating a specific disease or illness condition [8]. Further, consumers have turned their attention to plant sources for antioxidants instead of synthetic ones, which are feared to possess carcinogenic properties.
Several reports indicate that Cannabis species is a metal hyperaccumulator as it grows and has been used for soil bioremediation to remove contaminants [9,10]. Therefore, the safety of any hemp-derived product must be considered for heavy metal contamination. This implies that food products potentially made using hemp as an ingredient must be checked for safety before consumption.
Studies on the utilization of hemp-based products have suggested industrial and agricultural products such as the extraction of valuable chemical components, including fatty acids, cannabinoids, and terpenoids, as well as micro and nanofibers [11]. Furthermore, ongoing work is on its feasibility for feeding livestock and soil composting [12]. Little or no information exists presently on applying hemp by-products in food systems. Thus, this research aimed to assess the nutritional, safety (heavy metals), and functional (antioxidant properties and cannabinoid content) values of hemp flakes by comparing it with existing commercial hemp protein products. This work fits into the circular bio-economy [13], a proposed sustainability method for managing agro-resources, promising to be a better approach than our current harvest-use-and-dump approach. These research results could inform the food industry to incorporate hemp flakes as a safe, functional ingredient.
2Materials and methods
2.1Materials
A by-product obtained from a local facility expressing CBD oil from the Jinma variety's hemp buds and seeds was used. The dried material, hereafter referred to as ‘hemp flakes’ (H), was used in addition to three commercial hemp products (A, B, and C) purchased online (Fig. 1). The compared products were hemp protein powders selected from different geographical regions across the United States, Canada, and Italy with their specific names coded to prevent disclosing commercial products.
2.2Sample preparation
All samples were milled with a laboratory Waring blender (Dynamics Corporation of America, McConnelsburg, PA) and screened through a 2 mm mesh for proximate analytical determinations. All solvents used for extraction were obtained from Sigma-Aldrich (St. Louis, MO). Solvent ET–100 % ethanol (70 % v/v), solvent MET–100 % methanol (80 % v/v), solvent AQ–100 % distilled water, solvent EM–50 % methanol, 50 % ethanol) were used for extraction. For antioxidant assays with respective solvents, 5.0 ± 0.05 g of sample was added to 100 ml of the solvent and sonicated for 2 h, after which it was centrifuged (Thermo Fisher Scientific Sorvall Legend XTR, Waltham, MA, USA) at 3000×g at 4 °C for 20 min. The supernatant was filtered with Whatman #1 filter paper and evaporated to dryness using a rotary evaporator (Buchi Rotavapor R–215, New Castle, DE, USA). The dried extract was reconstituted to 10 ml by adding the respective solvent. All extracts were stored at 4 °C until further analysis.
2.3Proximate analysis
Proximate analysis of samples was conducted using the AACC-approved standard methods [14]. All chemicals were obtained from Sigma-Aldrich (St. Louis, MO). The air oven method was applied for moisture content determination (AACC44-15) using Sartorius (Practicum 224-1s, Wood Dale, IL, USA) weighing balance and a convection laboratory oven (Fisher Scientific Model # 6925, New Hampton, NH USA) at 130 °C for 60 min. For ash (AACC 08–01.01a), 3–5 g of sample was added to a pre-ignited, cooled, and weighed ashing porcelain crucibles and ignited at (600 ± 20 °C) overnight using a (Sybron Thermolyne model # 6020–12325, Dubuque, Iowa, USA). Crude protein (NX6.25) was determined by the Kjeldahl nitrogen method following AACC method 46–30.01. A Foss Tecator digestion system 2508 (Eden Prairie, MN, USA) and a Kjeltec auto 1030 analyzer (Waltham, MA, USA) were used. Crude fat was determined by following the AACC method 30–25.01 in a Foss Soxtec system 1043 H T (Eden Prairie, MN, USA). The crude fiber content (AOCS-approved method Ba 6a-05) of hemp samples was determined using an Ankom 2000 fiber analyzer (Ankom Technology, NY, USA). Approximately 0.95–1.00 g of milled hemp sample (passed through a 2 mm sieve) was placed in sealed filter bags and two blank bags for reference. The filter bags were then immersed in petroleum ether for 10 min to extract fat, drained, and placed in the analyzer's vessel with a bag suspender. The bags underwent digestion with 0.225 N H2SO4 and 0.313 N NaOH solutions, followed by rinsing with distilled water. After drying in an oven (102 ± 2 °C), the bags were ashed in a furnace (600 ± 15 °C) and weighed. The percentage of crude fiber was calculated by comparing the weight before and after ashing using Eq. (1). Carbohydrate content was determined by difference.Where W1 is the bag tare weight, W2 is the weight of the sample, W3 is the weight of organic matter, and C1 is the ash corrected factor.
2.4Antioxidant potential and free radical scavenging properties of hemp samples
2.4.1Total phenolic content
The Folin-Ciocalteau method was used to determine the total phenolic content in hemp samples as described by Singleton et al., [15]. Unknown samples were prepared by mixing 12.5 μl of the sample with 50 μl of distilled water. 12.5 μl of Folin-Ciocalteau reagent (Fisher Chemicals, Pittsburgh, PA, USA) was added to the mixture for 5 min. Finally, 125 μl of 7 % sodium carbonate anhydrous solution was added and shaken at room temperature for 90 min. Absorbance was read at a wavelength of 750 nm against a blank of distilled water using a microplate reader (Synergy HT, BioTek Instruments Inc. Winooski, Vermont, USA). The standard curve for determining total phenolic compounds was conducted with gallic acid (0–500 mg/l) (Fisher Chemicals, Pittsburgh, PA, USA), and the results were stated as mg GAE/100 g.
2.4.2Total flavonoids content
The aluminum chloride colorimetric method by Marinova et al. [16], was applied with little modifications for determining the total flavonoids in the samples. A standard curve of catechin (0.02–0.5 mg/l) (Cayman chemicals, Ann Arbor, MI, USA) was used, and the results were expressed as mg CE/100 g). Unknown samples were prepared by adding 125 μl of distilled water to 25 μl of extract in a well plate, followed by adding 7.5 μl of 5 % sodium nitrite and allowed to stand for 5 min 50 μl of 1M sodium hydroxide solution and 25.5 μl of distilled water were mixed briefly. The absorbance of the mixture was read against a blank distilled water at 510 nm with a microplate reader (Synergy HT, BioTek Instruments Inc. Winooski, Vermont, USA).
2.4.3Ferric reducing antioxidant potential (FRAP)
The ferric-reducing antioxidant potential of the samples was determined following the Benzie et al. [17], method with slight modifications. The FRAP reagent, freshly prepared, was prepared with 10 mmol 2,4,6-tri (2-pyridyl)-s-triazine (TPTZ) in 40 mmol hydrochloric acid, 20 mmol ferric chloride (FeCl3–6H20), and 300 mmol acetate buffer (pH 3.6). A standard curve was prepared using ferrous sulfate (0.1–1.0 mmol). Distilled water, 30 μl, and FRAP reagent, 30 μl, were added to 10 μl of the sample, allowed to mix briefly and read at an absorbance of 593 nm using a microplate reader (Synergy HT, BioTek Instruments Inc. Winooski, Vermont, USA).
2.4.41,1–diphenyl-1-picrylhydrazyl (DPPH) assay
A modified assay method described by Brand-Williams et al. [18], was applied to determine the radical scavenging ability of samples. DPPH solution, 0.1 mmol, was prepared in methanol, and 200 μl of the DPPH solution was added to 40 μl of sample in a well plate with the absorbance read at 517 nm using a plate reader (Synergy HT, BioTek Instruments Inc. Winooski, Vermont, USA). Results were expressed as a reduction in the percentage of DPPH using Eq. (2).where is the absorbance of the control and is the sample absorbance.
2.4.5Trolox equivalent antioxidant capacity (TEAC)
The Trolox equivalent antioxidant activity for samples was determined using the Miller et al. [19], method. A standard curve was first prepared with Trolox (0.02–0.80 mmol). 2,2-azino-bis-3-ethyl benzothiazoline-6-sulfonic acid (ABTS) was diluted with ethanol to obtain an absorbance of 0.70 ± 0.025 at a wavelength of 734 nm. The ABTS solution (7 mmol) was prepared and left overnight in the dark for the radical to develop. In the assay, 10 μl of sample and blank was combined with 290 μl of ABTS, followed by the absorbance measurement at 734 nm at 6 min using a microplate reader (Synergy HT, BioTek Instruments Inc. Winooski, Vermont, USA). The results were reported as micromole Trolox equivalent (μmol TE/100 g) of sample dry weight.
2.6Determination of cannabinoids by high-pressure liquid chromatography (HPLC)
Cannabinoids analysis of samples was conducted based on a modified method of Aubin et al., [21]. The analytical HPLC apparatus comprised a Varian Prostar LC system (USA) equipped with Prostar autosampler attached to a 100 μl loop and a Prostar 310 UV-VIS detector. Samples and standards were analyzed by reverse phase HPLC on a C18 column with 250 × 4.6 mm inner diameter and column particle size of 5 μm (Waters part #WAT054275, Ireland). The column heater was set to 25 °C to maintain a consistent temperature. DEA-exempt cannabidiol (CBD), cannabiodiolic acid (CBDA), cannabigerol (CBG), cannabichromene (CBC), and cannabinol (CBN) standards were obtained from Sigma-Aldrich Aldrich (St. Louis, MO, USA).
For cannabinoids analysis, an isocratic mobile phase consisting of A: HPLC-grade water acidified with 0.1 % trifluoroacetic acid and B:100 % HPLC-grade acetonitrile was employed at 20 %:80 % at a flow rate of 0.8 ml per min for separation and run for 30 min. Data was collected on a computer equipped with Varian Star Chromatography software – LC workstation version 6.41. UV spectra were recorded at 228 nm, allowing a 5-min re-equilibrium period within injections. Cannabinoid peak identification was conducted based on the standard peaks and was further employed for calculating the respective concentrations of all selected cannabinoids in the samples. Confirmatory tests were completed by spiking the samples with individual standards. The determination of concentrations of analytes was conducted in triplicate.
2.7Statistical analysis
Data was collected in triplicate, except ICP, which was completed in duplicate and reported as least square mean ± standard error of the means. Analysis of variance (ANOVA) was conducted at p ≤ 0.05, and the means were separated using Tukey's method. The statistical software used were SAS 9.4 (SAS Institute Inc, Cary, NC, 2020) for all ANOVA and means separation and Minitab 17.3.0 (Minitab Inc., 2016, PA, USA) for correlation analyses.
3Results and discussion
3.1Proximate analysis
Proximate composition of products depends on the species, geographical location of growth, growing season, and handling and processing conditions. The results of the proximate analysis of samples used in this work are shown in Table 1. The moisture content of the samples was within a narrow range of 6.0 and 7.0 %, and all samples were deemed dry and stable. Ash content ranged between 4.72 and 7.21 %, with hemp flakes (H) recording a significantly low amount of ash at 4.72 % (p ≤ 0.05). Crude fat content was highest (p ≤ 0.05) in H and could be attributed to the processing technique of adding a neutral oil to aid CBD oil extraction. Crude protein and carbohydrate content values for H were comparable to commercial hemp protein samples. Except for carbohydrate content, which was determined by difference, data on the chemical composition of moisture, ash, protein, and fat for all samples were similar to data reported by Pojic et al. [22], who worked on a similar by-product, a mechanically cold-pressed hemp meal. Hemp flake's crude fiber content (33.91 %) was within the range of the commercial hemp protein samples, which recorded crude fiber between 31.8 % and 42.4 %.Sample Moisture content (%) Ash (%) Crude fat (%) Crude protein (%) Crude fiber (%) Carbohydrates % H 7.73 ± 0.16a 4.72 ± 0.01c 15.38 ± 0.14a 31.62 ± 0.08b 33.91 ± 1.25c 6.64 ± 0.35c B 6.16 ± 0.84a 7.21 ± 0.30a 10.88 ± 1.11b 43.03 ± 0.18a 31.84 ± 1.49d 0.88 ± 0.66d A 7.74 ± 0.48a 5.47 ± 0.03b 10.09 ± 0.38b 22.00 ± 0.71d 42.24 ± 0.72a 12.46 ± 1.51a C 6.51 ± 0.48a 6.67 ± 0.23a 8.99 ± 0.02b 29.87 ± 0.42c 38.33 ± 0.64b 9.63 ± 0.61b
3.2Antioxidant and radical scavenging properties of hemp samples
3.2.1Total phenolic content
Total phenolic content measures the holistic content of all bioactive phytochemicals present in a product. Table 2 shows the total polyphenol content of hemp flakes and three commercial hemp protein products. The results ranged from a minimum of 1.40 mg GAE/100 g DW for hemp flake (H) from aqueous extraction to a maximum of 11.25 mg GAE/100 g DW in 1:1 ethanol/methanol extract for sample A. These results are similar to reported data when using similar extraction methods reported by Zago et al., [23]. The results for hemp flakes were similar for the ethanol (ET), methanol (MET), and 1:1 ethanol/methanol extracts compared to the commercial hemp protein products. The total polyphenol content in hemp flakes extracted from ethanol and 1:1 ethanol/methanol extracts were not significantly different (p ≤ 0.05) from those of sample B. The results suggest that the hemp flakes contained comparable or competitive total polyphenol content among the compared commercial protein samples. These results agree with [24] while investigating the effect of malting on hemp varieties' nutritional and antioxidant content. Azad et al. [25], also reported similar levels of hemp total polyphenol contents (7.30–15.40 mg GAE/100 g) while studying the impact of hot melt extrusion on hemp leaves for hemp decarboxylation of cannabidiolic acid into cannabidiol. The effect of solvent systems in the extraction of total polyphenols (Table 2) indicated ethanol extracts as the highest (p ≤ 0.05) in all samples except C and A, where 1:1 ethanol/methanol extracts produced significantly higher total polyphenol contents. These findings suggest that contrary to expectations, 1:1 ethanol/methanol solvent supersede ethanol or methanol extracts only with some samples.TPC mg GAE/g DW TFC mg CE/100 g DW Extraction solvent Extraction solvent Hemp AQ ET MET EM AQ ET MET EM H 1.40 ± 0.01c,D 2.53 ± 0.05a,C 2.23 ± 0.04b,D 2.48 ± 0.04a,C 0.04 ± 0.00d,B 0.30 ± 0.00a,B 0.11 ± 0.01c,C 0.17 ± 0.01b,D B 2.69 ± 0.03a,B 2.48 ± 0.06b,C 2.49 ± 0.01b,C 2.57 ± 0.02b,C 0.17 ± 0.01c,A 0.19 ± 0.01b,C 0.24 ± 0.01a,AB 0.23 ± 0.01a,C A 2.12 ± 0.05d,C 7.85 ± 0.05b,A 4.73 ± 0.05c,A 11.25 ± 0.01a,A 0.01 ± 0.00d,D 0.37 ± 0.00b,A 0.13 ± 0.00c,BC 0.93 ± 0.03a,A C 3.09 ± 0.05b,A 2.98 ± 0.01bc,B 2.88 ± 0.03c,B 3.33 ± 0.04a,B 0.02 ± 0.00b,C 0.30 ± 0.01a,B 0.30 ± 0.07a,A 0.29 ± 0.00a,B
3.2.2Total flavonoid content determination
The total flavonoid content of the hemp samples is presented in Table 2. Different solvents and samples yielded different flavonoid contents. The highest (p ≤ 0.05) flavonoid content (0.93 mg CE/100 g DW) was observed in sample A after extraction with 1:1 ethanol: methanol solvent. Aqueous extracts produced the lowest flavonoid content in all samples (p ≤ 0.05). The ethanol extract of flavonoids (0.30 mg CE/100 g DW) showed comparable (not significantly different p ≤ 0.05) amounts of flavonoids with C (0.30 mg CE/100 g DW) and higher when compared with B. The results indicate comparable properties to the flavonoid content of the commercial hemp protein products.
3.2.3DPPH free radical scavenging properties
In this experiment, antioxidant capacity was determined by the ability of the sample to scavenge the DPPH radical (1,1-diphenyl-2-picrylhydrazyl). DPPH radical is visually recognized by the decolorization in the cell plate during the experiment. A DPPH inhibition of more than 50 % is reported in the literature to signify a potent antioxidant [26]. The ethanol extract of sample H produced the highest (p ≤ 0.05) DPPH inhibition of 58.70 % among the hemp samples (Table 3). Aqueous and 1:1 ethanol/methanol extracts of H recorded DPPH inhibition rates similar to the commercial hemp protein samples. This observation suggests that the CBD by-product is a source of antioxidants and has similar powers to scavenge free radicals compared to commercial hemp protein products when processed similarly. Within H, it was observed that the ethanol extract produced the highest DPPH radical inhibition, followed by methanol and 1:1 ethanol/methanol solvent extraction. The observation here is that when ethanol and methanol are combined in a 1:1 ratio, the strength of the extraction for DPPH inhibition is lowered significantly (p ≤ 0.05) than using ethanol alone for the extraction.DPPH % TEAC (mmol TE/100 g) Extraction solvent Extraction solvent Hemp sample AQ ET MET EM AQ ET MET EM H 29.81 ± 0.65c,A 58.70 ± 0.87a,A 48.33 ± 0.23b,D 49.45 + 0.94b,C 55.83 ± 0.68c,C 121.03 ± 0.47a,A 121.28 ± 0.08a,A 107.94 ± 0.08b,A B 28.28 ± 1.21d,A 52.44 ± 0.15c,B 55.96 ± 0.23b,C 64.10 + 0.56a,A 87.14 ± 0.34c,B 120.86 ± 0.00a,A 120.86 ± 0.00a,AB 109.36 ± 1.46b,A A 22.97 ± 0.82c,B 52.10 ± 0.52b,B 57.75 ± 0.45a,B 50.47 + 0.70b,C 103.77 ± 4.50ab,A 121.03 ± 0.15a,A 121.37 ± 0.15a,A 97.42 ± 3.14b,A C 22.19 ± 0.56d,B 53.64 ± 0.34c,B 62.64 ± 0.23a,A 58.01 + 0.30b,B 98.66 ± 2.89b,A 121.03 ± 0.08a,A 120.18 ± 0.45a,B 102.91 ± 3.40b,A
3.2.4Trolox equivalent antioxidant activity (TEAC)
This Trolox equivalent assay determines the ability of antioxidants in a product to scavenge the 2,2′–azinobis (3-ethylbenzothiazoline-6-sulfonic acid) (ABTS) radical cation, with the extent of reaction visually captured as the bleaching of the blue-green color of ABTS [19]. The result of TEAC among hemp samples is shown in Table 3. It was generally high for all samples extracted with ethanol, methanol, or ethanol and methanol combination, ranging from 97.42 mmol TE/100 g in A extracted with EM to 121.86 mmol TE/100 g in B extracted with ethanol and methanol. Except for the aqueous extract, all hemp flake (H) extracts were among the highest (p ≤ 0.05) when compared with the commercial hemp samples. This result suggests that hemp flakes possess a parallel antioxidant capacity as commercial hemp products.
3.2.5Ferric reducing antioxidant power (FRAP)
The determination of FRAP of a biological material employs the reduction of ferric (Fe3+) to ferrous (Fe2+) by an antioxidant. The results of hemp samples in the present study are shown in Fig. 2. Hemp flake's aqueous extract recorded one of the least (p ≤ 0.05) FRAP, but when compared to the commercial varieties, it was not significantly different (p ≥ 0.05) from that of sample A aqueous extract. The ethanol extract for hemp flakes (H) recorded 68.00 ± 0.017 mmol Fe(II) and was intermediate among the results of corresponding commercial samples. Extracts from methanol and 1:1 ethanol/methanol recorded the highest FRAP values (p ≤ 0.05). Significant differences (p ≤ 0.05) were found in the antioxidant levels due to FRAP from the hemp samples. The decreasing order for FRAP recorded among the samples extracted with ethanol was A > B > H > C. These differences may be due to the varying sources of hemp samples used in this work. The results further indicate the impact of different solvents on the extraction of antioxidants. Ethanol extracts produced the highest FRAP, while aqueous extracts generally recorded the least significantly different (p ≤ 0.05) FRAP values. Different organic and aqueous solvent systems used singly or in multiple combinations of different ratios have been reported to impact antioxidant extracts significantly [27]. This is because the different solvent systems possess different polarities and can only dissolve antioxidants of similar polarities. Hemp flake's FRAP potential was superior to C and not statistically different from B with ethanol extraction, while aqueous extraction was similar to A. This indicates that hemp flakes have similar FRAP potential to commercial hemp products.
3.3Heavy metal content of hemp samples
The heavy metal contents in the hemp samples studied are presented in Fig. 3. Arsenic and lead, whose accumulation effects are among the most deleterious with implications for heart disease, high blood pressure, kidney, and bone disease [28], were not detected. Cadmium and cobalt recorded insignificant amounts in all the hemp samples; however, aluminum and copper were present in all samples. In decreasing order of content among samples, aluminum was A > H > C > B (p ≤ 0.05), while copper was A > C > H > B (p ≤ 0.05). This might be a characteristic behavior of industrial hemp since commercial samples were sourced from various locations, representing different geographical and growing conditions. The highest (p ≤ 0.05) heavy metal contamination in this work was in A (0.71± mg/kg DW) for aluminum. Establishing the level of heavy metals in hemp products is crucial, considering the nature of Cannabis species being used for phytoremediation of soils. Cannabis species are reportedly capable of extracting and bioaccumulating various metals from the soil to their stem, leaves, and branches compared to other species due to the possible presence of stress-tolerant genes [29]. For this reason, food product formulations constituted with hemp must be assessed to assure the safety of the consumer.
The results further indicate that all the levels of heavy metals found in all samples were within the permissible levels according to the Environmental Protection Agency (EPA) and the US Food and Drug Administration (FDA). The EPA has a permissible level of 10 ppm of arsenic in drinking water [30]. Copper and aluminum are not regarded as heavy metals because they have relatively low molecular weights; however, their accumulation in the body can pose risks to humans, such as Wilson's Disease. Copper is an essential mineral known to have critical functions in metabolism, first being able to take part in oxidation-reduction reactions as it moves from its cuprous (Cu+) to cupric forms (Cu2+). It also serves as a cofactor for many enzymes (cuproenzymes), including cytochrome c oxidase, whose action generates a proton gradient necessary for ATP synthesis, and lysyl oxidase, which initiates the formation of cross-links that stabilize elastin and collagen for the formation of strong connective tissue [31]. Despite all these essential attributes of copper, there could be an issue when it is overconsumed; even though it is rare in the general population, it is still essential to check the levels of copper in all foods.
Data comparison was conducted between the samples' chemical composition and heavy metal content (Table 4). Negative correlations were found between protein and copper (r = −0.832, p = 0.001) and protein and aluminum (r = −0.773, p = 0.003), as well as a positive correlation between fat and cobalt (r = 0.775, p = 0.003). These relationships, while correlation does not represent causation, and may need further research as we did not find similar reports in literature and more research on the topic would be needed to confirm the notion. If proven by further research, the results could be valuable information for hemp breeders aiming to develop high-protein or high-fat varieties.Protein Ash MC CFat Al Cu As Pb Co Ash 0.58
0.05MC −0.54
0.07−0.33
0.30CFat 0.15
0.64−0.65
0.02−0.14
0.66Al −0.77
0.003−0.64
0.010.64
0.030.17
0.59Cu −0.83
0.001−0.08
0.810.47
0.13−0.60
0.040.52
0.08As ** ** ** ** ** ** Pb ** ** ** ** ** ** ** Co 0.26
0.37−0.06
0.87−0.11
0.730.78
0.0030.15
0.64−0.53
0.08** ** Cd −0.004
0.99−0.54
0.07−0.06
0.870.64
0.030.05
0.88−0.41
0.18** ** 0.09
0.78
Overall, our results have provided evidence to posit that the oil by-product (hemp flakes) obtained from CBD oil extraction could potentially be regarded as a functional ingredient, as described by Ref. [32], “functional foods contain bioactive components with positive health effects reduce a compound with negative effects.” Further to the nutritional benefits obtained from the consumption, hemp flakes also possess bioactive compounds that confer systemic health benefits, such as reducing the risk of chronic disease or nullifying the effect of another component with a negative effect.
3.4Cannabinoid content in hemp samples
Hemp cannabinoids are known to be synthesized in acidic forms while hemp plants grow and, later, during harvest and processing operations, converted to their neutral forms by spontaneous heat or light-mediated reactions [4,33]. The calibration data and regression for five selected cannabinoids (cannabiodiolic acid–CBDA, cannabigerol–CBG, cannabidiol–CBD, cannabinol–CBN, and cannabichromene–CBC) and the results of cannabinoid content in hemp flakes and commercial hemp protein products are presented in Fig. 4, Fig. 5, and Table 5, respectively. Fig. 4 shows the regression coefficients of the standard analytes. The cannabinoids studied eluted in order of retention times as CBDA, CBG, CBD, CBN, and CBC (Fig. 5). Similar order of elution of these cannabinoids has been reported by several authors, including [21,34]. The results show variation in cannabinoids among the samples studied. This could be due to differences in hemp varietal and growing conditions since commercial samples were from various locations. All samples tested, including hemp flakes, contained CBDA and CBD. However, CBC was not detected in any of the samples, per the peaks and retention times during the HPLC (Fig. 5). CBN was only detected in hemp flakes (H) as was CBG only in sample C. Hemp flakes (H) showed the highest significantly different (p ≤ 0.05) amount of CBDA (0.144 mg/g DW) (Table 5) among all samples and cannabinoids studied. In comparison, sample C showed the highest significantly different (p ≤ 0.05) amount of CBD (0.574 mg/g).
The highest content of CBDA observed in hemp flakes could be attributed to the source of the hemp plant from where it was made (buds and flowers) versus the commercial hemp products, which are typically processed from hemp seeds. Hemp seeds have been commercially marketed for their nutrition, including essential fatty acids and high proteins, but not as a primary source of cannabinoids [35]. Samples A and C recorded the highest amounts of CBD, while hemp flakes recorded 0.038 mg/g DW. The heating process applied during the processing of hemp powders might have converted some CBDA into CBD. On the contrary, hemp flakes were obtained as a by-product of cold pressing for CBD oil extraction and thus might explain the low amounts of CBD and elevated levels of CBDA in hemp flakes since heat and light help the conversion of CBDA into its neutral form by decarboxylation into CBD [25,36].
Currently, in the United States, and depending on local regulations, the use of hemp ingredients and extracted CBD cannot be applied in foods on a federal basis [37]; however, some state/local regulations do allow them for recreational applications, including foods [38]. Cannabinoids, especially non-psychotic CBD, are reported to present functional benefits, including anti-inflammatory, anxiolytic, and neuroprotective properties, as well as being a muscle relaxant [3,4]. The by-product tested in this work was shown to contain similar kinds of cannabinoids and even higher amounts of specific cannabinoids compared to related commercial hemp food ingredients on the market and could, therefore, be recommended for use as a food ingredient to benefit consumers' health.
4Conclusions
It is predicted that by 2050, the world's population will reach 9 billion, and a sustained food supply will be a concern; therefore, it is appropriate to examine alternatives, including the exploration of agricultural waste materials. Hemp flakes as a by-product from CBD oil extraction could be utilized due to their nutritional and functional value.
The hemp flake used in this work demonstrated to hold nutritional and health components comparable to related commercial products. The antioxidant levels showed variations attributed to the source of hemp material and solvent extraction method. Hemp flakes exhibited high and similar antioxidant properties as measured by TPC, TFC, FRAP, and TEAC and possessed comparable radical scavenging properties as measured by DPPH. The hemp by-product showed comparative amounts of cannabinoids with the highest content of cannabidiolic acid, which is known to break down to cannabidiol and possess functional benefits. Further, the results of this work have exemplified that hemp flakes generated from CBD oil extraction have a considerable nutritional and functional value that supports its potential to be incorporated in food preparations as an ingredient. It was also established that the hemp flakes contained levels below the permissible levels of heavy metals in foods, according to health and environmental agencies. It is concluded that the by-product from CBD oil extraction could be utilized as an ingredient in food processing, such as a composite with other ingredients to complement nutrition and health functionality for consumers.
Ethics statement
This work did not use animal or human subjects to collect data.
Data availability
Data supporting this work's findings have not been deposited in any repository and would be available upon reasonable request.
Declaration of competing interest
The authors declare that they have no known competing personal or financial relationships that could have appeared to influence the work in this article.
Acknowledgment
We acknowledge funding from the 10.13039/100000199USDA Evans Allen Program (ALAX - 012-1422 EA) and 10.13039/100024174Alabama Agricultural & Mechanical University for this project. We acknowledge Dr. Lloyd Walker for allowing us to work in the Chemistry Laboratory (HPLC) in the Agricultural Research Center and Mr. Simon Ogutu of the Food and Animal Sciences Department, AAMU, for providing technical laboratory support.