Tetrahydrocannabinol
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Tetrahydrocannabinol (THC), in full (−)-trans-Δ9-tetrahydrocannabinol, is the cannabinoid that mainly produces the psychotropic effects of cannabis.[1] The living plant holds little of it. The glandular trichomes of the female flowers make its carboxylic acid, tetrahydrocannabinolic acid (THCA). THC forms from the acid without the help of an enzyme as the harvested material dries, is stored or is smoked.[2][3][4]
The balance between THC and cannabidiol (CBD) in a plant is inherited almost as a single gene and defines its chemotype. It differs widely between the landraces of different regions. In a study of 157 accessions grown together, the allele for THC production was fixed in the narrow-leaflet drug landraces and rare in the hemp and ruderal populations of Europe and Central Asia.[4]
The structure of THC was established in 1964 by Y. Gaoni and Raphael Mechoulam, who isolated it from hashish.[5][6] THC content in the dried flowering tops of female plants is the measure by which many countries separate hemp from drug cannabis, at a threshold of 0.3 per cent taken from a taxonomy of the genus published in 1976.[7]: 82
Formation in the plant
Cannabinoids are made mainly in the glandular trichomes that develop on female flowers and to a lesser extent on leaves.[2] In figures compiled for the 2009 laboratory manual of the United Nations Office on Drugs and Crime (UNODC), total THC is 10–12 per cent in female flowers, 1–2 per cent in leaves, 0.1–0.3 per cent in stalks and below 0.03 per cent in roots.[8]: 14 The bracteole that encloses the seed carries the densest glandular trichomes on the plant and so the highest THC concentration, but the seed itself contains no THC; THC found in hemp seed or seed oil comes from plant material left with the seed.[8]: 19
Cannabinoids are prenylated polyketides, built from fatty-acid and isoprenoid precursors. A polyketide synthase joins hexanoyl-CoA to three molecules of malonyl-CoA. A second enzyme, olivetolic acid cyclase, closes the chain into olivetolic acid. Olivetolic acid takes on a geranyl group to become cannabigerolic acid (CBGA), which oxidocyclase enzymes convert into THCA or cannabidiolic acid (CBDA).[2] The enzyme of the THC branch, THCA synthase, closes a ring in the geranyl part of CBGA. It carries a bound flavin cofactor, uses molecular oxygen and releases hydrogen peroxide. Its gene, cloned in 2004 from the leaves of a Mexican strain, was the first gene of cannabinoid biosynthesis to be cloned. Its sequence shows no similarity to the enzymes that cyclise monoterpenes and is closest to the berberine bridge enzyme of the California poppy, an enzyme of alkaloid biosynthesis. THCA had earlier been thought to form from CBDA; the work that identified the synthase showed that it is made directly from CBGA.[3][8]: 14
A genetic and physical map made from a cross between the drug strain Purple Kush and the hemp variety Finola placed the THCA synthase and CBDA synthase genes at closely linked loci, each inside a region of more than 250 kilobases rich in retrotransposons and largely unlike its counterpart in the other parent. Both regions are embedded in about 40 megabases of repetitive DNA in which recombination is rare. A third gene of the pathway, the aromatic prenyltransferase that makes CBGA, is tightly linked to a known marker for total cannabinoid content.[9] A pangenome of 193 genomes published in 2025 found very little diversity in the cannabinoid synthase genes themselves, although the region around them is highly variable and holds several pseudogene copies.[10]
From acid to THC
Fresh plant material holds THC only at low levels. Decarboxylation, the loss of the acid's carboxyl group as carbon dioxide, takes place during drying, storage and heating, smoking included.[3][4][8]: 32 When cannabis extracts were heated in the laboratory at temperatures between 80 and 145 °C, THCA lost its carboxyl group in a first-order or pseudo-first-order reaction about twice as fast as CBDA or CBGA. The reaction gave THC with no side reactions or by-products.[11]
Gas and liquid chromatography are the techniques most often used to measure cannabinoids. The acids cannot be analysed by gas chromatography without being decarboxylated or converted to a derivative first.[11] A gas chromatograph run on an underivatised extract therefore reports "total THC", the free THC plus the THC formed from THCA. In potency testing a laboratory can measure THCA and THC separately or measure total THC. National law sometimes makes the choice. Where it does not, the UNODC manual calls measuring total THC common practice, since that figure comes closest to the pharmacological activity of the material. Some injectors decarboxylate THCA completely and others poorly. A high injection temperature can also break THC down.[8]: 32–33, 39
After drying, THC is still oxidised by air and ultraviolet light to cannabinol (CBN), a degradation product that does not occur in the living plant.[8]: 14, 26 Heating in the presence of oxygen and light has the same effect.[11] Freshly and carefully dried material contains no CBN. The ratio of CBN to THC can be used to estimate a sample's age: one study put samples with a ratio below 0.013 at less than six months old and those with a ratio between 0.04 and 0.08 at one to two years.[8]: 19–20
Structure and naming
THC has the formula C21H30O2 and a molecular weight of 314.46. It is a viscous oil, insoluble in water (2.8 mg per litre at 23 °C) and soluble in ethanol, chloroform and hexane. THCA, C22H30O4, has a molecular weight of 358 and decarboxylates at about 125–150 °C.[8]: 21 Like the other cannabinoids, THC is a terpenophenolic compound, built from an alkylresorcinol and a monoterpene unit.[4][3] The plant makes the (−)-enantiomer.[6]
Two numbering systems are in use for the ring positions. In the formal pyran numbering, now the more common, the compound is Δ9-THC. Older literature uses a monoterpenoid numbering in which the same compound is Δ1-THC. Its isomer Δ8-THC is Δ6-THC in that system.[6] The 2004 study that cloned the synthase uses the older form, Δ1-tetrahydrocannabinolic acid.[3]
The plant also makes a homologue of THC with a three-carbon side chain in place of five, tetrahydrocannabivarin (THCV), in a parallel series that starts from cannabigerovarin.[4]
Variation between plants
How much THC a plant carries depends on its sex and maturity, the day length, temperature, nutrients, ultraviolet light and the part of the plant sampled. In the mature unpollinated inflorescence of a female plant the THC or CBD content can exceed 10 per cent of dry weight, against under 1 per cent in the primary leaves. The ratio of THC to CBD, by contrast, is under simple genetic control.[4]
Chemotypes
In 1973 Ernest Small and Beckstead divided plants into three chemotypes by the ratio of THC to CBD: high (chemotype I), intermediate (II) and low (III). In plants from 96 accessions measured by Karl W. Hillig and Paul Mahlberg, the ratios fell into three discrete groups: more than 50 for chemotype I, 0.25 to 1.50 for II and less than 0.20 for III. A plant's chemotype is set at a young age and stays stable beyond the seedling stage. THC was found in every plant they analysed, chemotype III included.[4]
In 2003 Etienne de Meijer and colleagues proposed that the THCA and CBDA synthases are encoded at a single locus by two alleles, BT and BD, a model supported by the roughly 1:2:1 ratio of the three chemotypes in crosses between drug and hemp plants.[4][9] The 2019 genome map revised it: the two genes are not alleles of one locus but sit at separate, closely linked loci. No copy of the THCA synthase gene, working or disabled, was found in the hemp parent.[9]
Landraces and geography
The cannabinoid content of a strain is associated with its geographic origin. Most of Small and Beckstead's high-THC strains came from latitudes south of 30° N.[4] Hillig and Mahlberg grew 157 accessions of known origin together at Indiana University and scored their THC and CBD alleles. In their two-species scheme the frequency of the THC allele ran from 0.04 in the ruderal accessions of Central Asia to 1.00 in the narrow-leaflet drug landraces. It was significantly higher in every group they placed in Cannabis indica than in those they placed in Cannabis sativa. Nearly all accessions with a low frequency came from north of 35° N, but so did several with a high one, among them drug landraces from Afghanistan and Pakistan and hemp landraces from China, Japan and South Korea.[4]
The southern and eastern Asian hemp landraces in that study were mostly chemotype I plants that made less THC than the drug landraces. Hillig and Mahlberg explained the half-century struggle of European breeders to lower the THC content of Chinese hemp partly by the breeders' failure to recognise that East Asian hemp belongs to the C. indica gene pool. They found the THC allele at high frequency in feral populations from the north-western Himalayas too and concluded that it was common in that gene pool before domestication. On their reading, human selection raised it to fixation only in the narrow-leaflet drug landraces, which are grown for their flowers, each plant's harvest used on its own. The wide-leaflet drug landraces were traditionally grown for hashish, a pooled product of many plants whose individual chemotypes the grower usually does not know.[4]
John McPartland and Small, classifying the drug landraces and their wild relatives in 2020, separate the South Asian from the Central Asian domesticates partly by their THC to CBD ratio, at least 7 in the first and below 7 in the second.[7]: 81 They summarised published THC figures for each of their four varieties of Cannabis sativa subsp. indica:[7]: 95–101
| Variety | Origin and status | Weighted mean THC | Range or highest value |
|---|---|---|---|
| var. indica | South Asian domesticate | 3.97% | up to 12.5% |
| var. himalayensis | South Asian wild-type | 1.49% | 0.06–9.3% |
| var. afghanica | Central Asian domesticate | 5.69% | up to 14.5% |
| var. asperrima | Central Asian wild-type | 1.49% | 0.4–4.47% |
They relate the high combined THC and CBD of var. afghanica, the highest of the four, to its dense covering of glandular trichomes.[7]: 98 THCV is common in landraces from South Asia and Africa.[7]: 95 McPartland and Small regard the Central and South Asian landraces as corrupted by foreign germplasm brought into their centres of diversity. They see the worldwide introgressive hybridisation of "Indica" and "Sativa" as a threat to the crop's agrobiodiversity.[7]: 85–86
The 0.3 per cent line
Small and Arthur Cronquist separated Cannabis sativa subsp. sativa from subsp. indica in 1976 by THC content: below 0.3 per cent in the dried flowering tops of female plants for the first, at least 0.3 per cent for the second. Many countries have since written the 0.3 per cent figure into the rules that separate fibre-type from drug-type plants.[7]: 82 In 2009 the upper legal limit for industrial hemp was 0.2 per cent in Europe and 0.3 per cent in Canada. The UNODC manual also describes a ratio test: if the summed gas-chromatograph peak areas of THC and CBN exceed that of CBD, the material is drug-type.[8]: 20
About a fifth of Hillig and Mahlberg's chemotype III plants, the CBD-dominant class, had more than 0.3 per cent THC. They argued that a limit of 0.8 per cent would fit the natural range of such plants better.[4]
Breeding for THC
In illicit cannabis seized in the United States and analysed at the University of Mississippi, the THC content of plant material rose from about 4 per cent in 1995 to about 12 per cent in 2014. CBD fell over the same years, from about 0.28 per cent in 2001 to under 0.15 per cent in 2014. The ratio of THC to CBD went from about 14 to about 80. Mahmoud A. ElSohly and his colleagues read the changing ratio as a sign that producers were breeding plants for higher THC. They also recorded a shift in production from ordinary marijuana to sinsemilla.[12] In Europe the mean THC content of herbal cannabis rose from 5.00 to 10.22 per cent between 2006 and 2016. Resin rose from 8.14 to 17.22 per cent. Tom Freeman and his colleagues considered the replacement of the traditional kif landrace in Morocco by high-potency strains a convincing explanation for the change in resin.[13]
Isolation and identification
The first cannabinoid to be isolated was cannabinol, at the end of the 19th century.[6] In 1896 the chemists T. Barlow Wood, W. T. Newton Spivey and Thomas Hill Easterfield analysed charas from the North-Western Provinces of India and gave the name cannabinol to a red oil they separated from it.[14] R. S. Cahn worked out the structure of cannabinol in the early 1930s. In 1940 the laboratories of R. Adams in the United States and Lord Todd in Britain synthesised it. THC was first extracted in 1942 by Wollner, Matchett, Levine and Loewe, most likely as a mixture of the Δ8 and Δ9 isomers.[6]
Gaoni and Mechoulam isolated Δ9-THC from cannabis in 1964 and published its structure under the title "Isolation, structure, and partial synthesis of an active constituent of hashish".[5][6] The racemic compound was first synthesised in Mechoulam's laboratory in 1965.[6] Research in the 1960s and early 1970s established that THC produces most of the psychotropic effects of cannabis.[1]
Action at cannabinoid receptors
In 1975 David Lawrence and Edward Gill proposed that cannabinoids act by disordering the lipids of cell membranes, with no need for a specific receptor. In 1988 Allyn Howlett's laboratory, with Bill Devane, found high-affinity binding sites for a labelled synthetic cannabinoid in rat brain. The rat CB1 receptor was cloned in 1990 and a second receptor, CB2, in 1993. In 1992 Devane, by then working in Jerusalem with Mechoulam, isolated from pig brain the first of the body's own cannabinoids, named anandamide from the Sanskrit ananda, bliss.[6]
CB1 receptors are found mainly on nerve cells and CB2 receptors mainly on cells of the immune system. THC is a partial agonist at both. It binds CB1 about as strongly as anandamide does but activates it less. Its activation of CB2 is weaker still. In some experiments it has acted as a CB1 antagonist, which Roger Pertwee relates to the low density or weak coupling of the receptors in some neurons. THC entered clinical use in the 1980s under the name dronabinol, as did its synthetic analogue nabilone.[1]
See also
References
- ↑ 1.0 1.1 1.2 Pertwee, R.G. (2008). "The diverse CB1 and CB2 receptor pharmacology of three plant cannabinoids: Δ9-tetrahydrocannabinol, cannabidiol and Δ9-tetrahydrocannabivarin". British Journal of Pharmacology. 153 (2): 199–215. doi:10.1038/sj.bjp.0707442. PMC 2219532.
- ↑ 2.0 2.1 2.2 Gagne, S.J.; Stout, J.M.; Liu, E.; Boubakir, Z.; Clark, S.M.; Page, J.E. (2012). "Identification of olivetolic acid cyclase from Cannabis sativa reveals a unique catalytic route to plant polyketides". Proceedings of the National Academy of Sciences. 109 (31): 12811–12816. doi:10.1073/pnas.1200330109. PMC 3411943.
- ↑ 3.0 3.1 3.2 3.3 3.4 Sirikantaramas, S.; Morimoto, S.; Shoyama, Y.; Ishikawa, Y.; Wada, Y.; Shoyama, Y.; Taura, F. (2004). "The gene controlling marijuana psychoactivity: molecular cloning and heterologous expression of Δ1-tetrahydrocannabinolic acid synthase from Cannabis sativa L." Journal of Biological Chemistry. 279 (38): 39767–39774. doi:10.1074/jbc.M403693200.
- ↑ 4.00 4.01 4.02 4.03 4.04 4.05 4.06 4.07 4.08 4.09 4.10 4.11 Hillig, K.W.; Mahlberg, P.G. (2004). "A chemotaxonomic analysis of cannabinoid variation in Cannabis (Cannabaceae)". American Journal of Botany. 91 (6): 966–975. doi:10.3732/ajb.91.6.966.
- ↑ 5.0 5.1 Gaoni, Y.; Mechoulam, R. (1964). "Isolation, structure, and partial synthesis of an active constituent of hashish". Journal of the American Chemical Society. 86 (8): 1646–1647. doi:10.1021/ja01062a046.
- ↑ 6.0 6.1 6.2 6.3 6.4 6.5 6.6 6.7 Pertwee, R.G. (2006). "Cannabinoid pharmacology: the first 66 years". British Journal of Pharmacology. 147 (S1): S163–S171. doi:10.1038/sj.bjp.0706406. PMC 1760722.
- ↑ 7.0 7.1 7.2 7.3 7.4 7.5 7.6 McPartland, J.M.; Small, E. (2020). "A classification of endangered high-THC cannabis (Cannabis sativa subsp. indica) domesticates and their wild relatives". PhytoKeys. 144: 81–112. doi:10.3897/phytokeys.144.46700.
- ↑ 8.0 8.1 8.2 8.3 8.4 8.5 8.6 8.7 8.8 United Nations Office on Drugs and Crime (2009). Recommended Methods for the Identification and Analysis of Cannabis and Cannabis Products (PDF). Manual for use by national drug analysis laboratories (Revised and updated ed.). New York: United Nations. ISBN 978-92-1-148242-3. ST/NAR/40.
- ↑ 9.0 9.1 9.2 Laverty, K.U.; Stout, J.M.; Sullivan, M.J.; Shah, H.; Gill, N.; Holbrook, L.; Deikus, G.; Sebra, R.; Hughes, T.R.; Page, J.E.; van Bakel, H. (2019). "A physical and genetic map of Cannabis sativa identifies extensive rearrangements at the THC/CBD acid synthase loci". Genome Research. 29 (1): 146–156. doi:10.1101/gr.242594.118. PMC 6314170.
- ↑ Lynch, R.C.; Padgitt-Cobb, L.K.; Garfinkel, A.R.; Knaus, B.J.; Hartwick, N.T.; et al. (2025). "Domesticated cannabinoid synthases amid a wild mosaic cannabis pangenome". Nature. 643 (8073): 1001–1010. doi:10.1038/s41586-025-09065-0. PMC 12286863.
- ↑ 11.0 11.1 11.2 Wang, M.; Wang, Y.-H.; Avula, B.; Radwan, M.M.; Wanas, A.S.; van Antwerp, J.; Parcher, J.F.; ElSohly, M.A.; Khan, I.A. (2016). "Decarboxylation study of acidic cannabinoids: a novel approach using ultra-high-performance supercritical fluid chromatography/photodiode array-mass spectrometry". Cannabis and Cannabinoid Research. 1 (1): 262–271. doi:10.1089/can.2016.0020. PMC 5549281.
- ↑ ElSohly, Mahmoud A.; Mehmedic, Zlatko; Foster, Susan; Gon, Chandrani; Chandra, Suman; Church, James C. (April 2016). "Changes in cannabis potency over the last 2 decades (1995–2014): analysis of current data in the United States". Biological Psychiatry. 79 (7): 613–619. doi:10.1016/j.biopsych.2016.01.004. PMID 26903403.
- ↑ Freeman, Tom P.; Groshkova, Teodora; Cunningham, Andrew; Sedefov, Roumen; Griffiths, Paul; Lynskey, Michael T. (2019). "Increasing potency and price of cannabis in Europe, 2006–16". Addiction. 114 (6): 1015–1023. doi:10.1111/add.14525. PMC 6590252.
- ↑ Wood, T. Barlow; Spivey, W. T. Newton; Easterfield, Thomas Hill (1896). "Charas. The resin of Indian hemp". Journal of the Chemical Society, Transactions. 69: 539–546. doi:10.1039/ct8966900539 – via Zenodo.