Tall oil, sodium salt

Endpoint summary

Description of key information

Tall Oil Soap is a UVCB substance (Substance of Unknown or Variable composition, Complex reaction products or Biological materials) obtained from naturally- occurring compounds extracted from tree species, principally of the genera Pinus, Picea, Betula and Populus. The substance is obtained from the Kraft pulping process used in the EU to make paper. Additionally, it is variable in composition and its precise composition depends both on the process conditions used to extract it, the time of year when it is extracted, and the species of trees used.  Consequently, the material contains a great number of chemical compounds, and at least 80 have been identified, although specific constituents and their relative proportions vary from batch to batch. The constituents include fatty acids, rosin acids (abietic acid and its derivatives), neutral compounds and polymers such as lignin and cellulose fibres. The neutrals include phytosterols, terpenoids and rosin alcohols; examples of phytosterols are β-sitosterol, stigmasterol and campesterol.

Crude Tall Oil (CTO) is produced from the acidulation of Tall Oil Soap (TOS) and as such the constituents of both substances are similar but in different proportion. In TOS the fatty acids and rosin acids are present as their sodium salts.

Additional information

TOS is a UVCB substance containing at least 80 different constituents. No in vitro or in vivo toxicokinetic studies are available for the whole product and it would not be scientifically feasible to conduct such a test.

The constituents of TOS are all naturally occurring secondary metabolites of pine trees and many other types of plant. Fatty acid constituents in particular are an important dietary source for humans, and the sterol constituent β-sitosterol is used as a supplement in certain types of fat spreads due to its cholesterol-lowering properties.

The properties of the substance are assessed using the 'block approach'. The key physico-chemical properties of each 'block' have been determined based on the weighted average of each constituent in the block. The assessment of toxicokinetic behaviour for Block 3 (Water) and Block 12 (Lignin, cellulose fibre and oligomeric acids) is not relevant being a high molecular weight compounds with uptakes being unlikely.

The following summary has therefore been prepared based on the predicted physicochemical properties of the constituents of the substance. The data have been used in algorithms which are the basis of many computer-based physiologically based pharmacokinetic or toxicokinetic (PBTK) prediction models. Although these algorithms provide quantitative outputs, for the purposes of this summary only qualitative statements or predictions will be made because of the remaining uncertainties that are characteristic of prediction models.

The main input variable for the majority of the algorithms is the log K_ow. By using this and where appropriate, other known or predicted physicochemical properties of constituents of TOS reasonable predictions or statements may be made about its potential absorption, distribution, metabolism and excretion (ADME) properties.

Relevant human exposure can occur via the oral or dermal routes

Absorption and distribution

Some of the existing in vivo toxicity studies for constituents of TOS, for example the repeated oral exposure with Distilled Tall Oil (Clubb and Daly, 2002), show clear evidence of absorption and distribution in vivo (particularly to the liver). Uptake via micellular solubilisation is an important mechanism for substances with log K_ow >4 and low water solubility (REACH Technical Guidance Chapter R7c).

In the only available dermal toxicity study, read-across from the related substance Crude Tall Oil (Bernat, 2005), there were no clinical signs of toxicity or gross pathological findings at the limit dose level of 2000 mg/kg/day. It is therefore not possible to determine if any dermal absorption occurred. Dermal absorption is optimal for substances with log K_ow 2-3 and is favourable for those with log K_ow in the range 0-4. All the constituents of TOS have log K_ow >4 at pH 5.5 (relevant for dermal exposure) and therefore dermal uptake may be limited. For those constituents in the log K_ow range 4-6 (relevant for the rosin acids, fatty acids up to C16 and neutral constituents in Block 4), the rate of penetration may be limited by the rate of transfer between the stratum corneum and the epidermis, but uptake into the stratum corneum will be high. Above a log K_ow of 6 (relevant for fatty acids above C16 as well as the neutral constituents excluding Block 4), the rate of transfer between the stratum corneum and the epidermis will be slow and will limit absorption across the skin. Uptake into the stratum corneum itself may be slow. (REACH Technical Guidance Chapter R7c). Dermal absorption could be enhanced by the pH-induced corrosive effects of TOS.

The positive Local Lymph Node Assay skin sensitisation test for CTO (Weber, 2005), which is read-across to TOS, indicates that some constituents do penetrate the epidermis and reach the lymph nodes via epidermal Langerhans cells.

Significant inhalation exposure is not expected for this non-volatile substance, and no toxicity data are available for this exposure route.

Fatty acids are required components in the diet of animals and are metabolised in cells to generate cellular energy in the form of adenosine triphosphate (ATP), and as such, they are the major source of metabolic energy in the western diet.  Additionally, dietary deficiencies of certain unsaturated fatty acids (essential fatty acids, which cannot be synthesised in vivo but must be obtained via the diet) such as linoleic acid (18:2 omega-6) and linolenic acid (18:3 omega-3) during human development has been shown to result in reduced levels of 20:4 omega-6 and 22:6 omega-3 fatty acids in the developing central nervous system, and this has been associated with altered learning behaviour and visual function (Innis, 1993). This is due to their function as major acyl components of cell membrane phospholipids in the central nervous system. Consequently, fatty acids may be considered as both sources of energy-rich food and essential components of the animal body.  In the normal diet, fatty acids are mainly ingested as their triglyceride esters (fats) and are broken down in the small intestine (mainly the duodenum) where they are hydrolysed to mono- and di-glycerides plus free fatty acids by pancreatic lipase.  Once adsorbed, free fatty acids may either be used to make triacylglycerol, which is stored as fat as a potential energy source or broken down and catabolised to CO₂ via oxidative metabolism as an instant source of energy.  The catabolism of a saturated fatty acid such as stearic acid is rather different from that for unsaturated fatty acids (linoleic, oleic and bovinic) and will be considered separately.

Primary digestion of fats takes place in the stomach under the influence of salivary and gastric lipases which primary catalyse hydrolysis in the 1-position. In the duodenum, fats are emulsified by bile acids, which act as a detergent, synthesised in the hepatic gall bladder and which are then transported via the bile duct to the duodenum.  These emulsified fats then become substrates for pancreatic lipases, which enter the duodenum from the pancreatic duct. Pancreatic lipase completes the hydrolysis to fatty acids by catalysing hydrolysis in the 2- and 3-positions. The resultant fatty acids then form mixed micelles with the bile acids and form part of the fraction called chylomicrons. The micelles function as a transport vehicle to deliver the fatty acids to the apical membrane of enterocytes, which are cells lining the small intestine that adsorb and transport small molecules from the lumen.  

The mixed micelles in the small intestinal lumen promote the absorption of fatty acids by facilitating transport of these lipids across the water layer adjacent to the surface of the apical membrane of enterocytes. The micelle particle itself does not penetrate the cell membrane, but it facilitates passage across a diffusion barrier that is located at the intestinal lumen-membrane interface for the uptake by the enterocytes. Mucous coating the intestinal mucosa is also a diffusion-limiting barrier, as cholesterol molecules may be extensively bound to surface mucins prior to transfer into the enterocyte. Presence of the mucins are necessary for normal intestinal uptake and absorption of cholesterol in mice, but it appears not to effect fatty acid uptake (Wang et al, 2004).

There are three putative pathways for the uptake of fatty acids and their transport across the apical membranes of enterocytes:

- Fatty acid translocase (CD36), alone or together with the peripheral membrane-bound fatty acid-binding protein (FABPpc) binds medium-chain and long-chain fatty acids at the cell surface to increase their local concentrations. This helps CD36 actively transport fatty acids across the apical membrane of the enterocyte. Once at the inner side of the membrane, these fatty acids are bound by cytoplasmic FABP (FABPc) before entering metabolic pathways.

- Medium-chain and long chain fatty acids are transported by fatty acid transport protein 4 (FATP4). These fatty acids may be rapidly activated by plasma membrane acyl-CoA synthetase 1 (ACS1) to form acyl-CoA esters.

- Short-chain fatty acids may traverse the apical membrane by simple passive diffusion and may be absorbed into the mesenteric venous blood and then the portal vein, however, this mechanism is not relevant to the fatty acids present in TOS.

Fatty acids that escape intestinal absorption are excreted in the faeces.

Absorption

Oral

When oral exposure takes place it can be assumed, except for the most extreme of insoluble substances, that uptake through intestinal walls into the blood occurs. Uptake from intestines can be assumed to be possible for all substances that have appreciable solubility in water or lipid. Other mechanisms by which substances can be absorbed in the gastrointestinal tract include the passage of small water-soluble molecules (molecular weight up to around 200 g/mol) through aqueous pores or carriage of such molecules across membranes with the bulk passage of water (Renwick, 1993).

With the exception of Block 4, the mean molecular weight of constituents in Blocks 1-2 and 5-11 are above to the favourable range for absorption. Additionally, the constituents have low water solubility, making systemic exposure limited.

Constituents in Block 4 Terpenes are slightly soluble with a representative water solubility of 5.7 mg/L at 20°C and have a molecular weight favourable for absorption, therefore should oral exposure occur it is possible that systemic exposure may also occur.

Following a repeated dose of phytosterol esters, no systemic effects have been observed. However, when the test substance was rosin, systemic effects were observed in both female and male rats based on altered body weight and weight gain following exposure at 7500 ppm.

Dermal

The fat solubility and therefore potential dermal penetration of a substance can be estimated by using the water solubility and log K_ow values. Substances with log K_ow values between 1 and 4 favour dermal absorption (values between 2 and 3 are optimal) particularly if water solubility is high.

Constituents in Blocks 1 and 4-11 have a log K_ow>4 and have low to moderate solubility. Therefore, dermal absorption of these constituents is unlikely to occur, since they are not sufficiently soluble in water to partition from the stratum corneum into the epidermis. However, dermal absorption could be increased if damage to the skin occurs due to corrosive effects of the substance.

For constituents in Block 2 Rosin Acid Sodium Salts, the water solubility and log K_ow are in the favourable range. For these constituents, absorption across the skin may occur as the substance is likely to be sufficiently hydrophilic to cross the lipid-rich environment of the stratum corneum.

No clinical signs of toxicity, no deaths and no treatment related effects on body weight were observed when an acute dermal application of CTO was applied. Similarly, no effects were observed following an acute dermal application of plant sterols. Nevertheless, CTO was regarded as a skin sensitiser. No studies are available for repeated dermal dose toxicity.

Inhalation

There is a Quantitative Structure-Property Relationship (QSPR) to estimate the blood:air partition coefficient for human subjects as published by Meulenberg and Vijverberg (2000). The resulting algorithm uses the dimensionless Henry coefficient and the octanol:air partition coefficient (K_oct:air) as independent variables.

 

Using these values foreach of the Blocks predicts an extremely low blood:air partition coefficient (<1E-06:1) meaning that if lung exposure occurs, uptake into the systemic circulation is not likely.

There are no inhalation toxicity studies to confirm evidence of absorption of the test material.

Distribution

For blood:tissue partitioning a QSPR algorithm has been developed by De Jongh et al. (1997) in which the distribution of compounds between blood and human body tissues as a function of water and lipid content of tissues and then-octanol:water partition coefficient (K_ow) is described.

Using the representative log K_ow value for each of the constituent ‘blocks’the algorithm predicts that, should systemic exposure occur, the distribution into the main body compartments is similar for all constituents, as follows: fat >> brain > liver ≈ kidney > muscle

Table 5.1.1 Tissue:blood partition coefficients

	Log Kow	Kow	Liver	Muscle	Fat	Brain	Kidney
BLOCK 1: Sodium salts of saturated and unsaturated C14-C20 fatty acids	5.16	1.45E+05	8.9	5.4	113.8	14.2	7.3
BLOCK 2: Rosin Acid Sodium Salts	3.7	5.01E+03	7.9	4.9	111.5	6.6	4.4
BLOCK 4: Terpenes	4.27	1.86E+04	8.5	5.3	141.6	10.5	6.3
BLOCK 5: Sesquiterpenes	6.27	1.86E+06	8.9	5.5	113.9	18.2	8.1
BLOCK 6: Abietenes and labdanes	8.04	1.10E+08	8.9	5.5	113.9	20.3	8.4
BLOCK 7: C30 Branched alkenes	10	1.00E+10	8.9	5.5	113.9	20.6	8.4
BLOCK 8: 3,5-Dimethoxystilbene	4.69	4.90E+04	8.8	5.4	113.7	11.8	6.6
BLOCK 9: Rosin Alcohol and aldehydes isomers	6.22	1.66E+06	8.9	5.5	113.9	18.1	8.1
BLOCK 10: C20-C35 alcohols and terpene alcohols.	9.02	1.05E+09	8.9	5.5	113.9	20.5	8.4
BLOCK 11: Sterols	9.42	2.63E+09	8.9	5.5	113.9	20.6	8.4

 

Sodium ions will enter the body's natural homeostatic processes.

Metabolism and excretion

Available published information on the metabolism and other biological behaviour of constituents of TOS has been reviewed and is summarised below. The metabolic pathways is attached in Section 13.

Block 1: Saturated and unsaturated C16-C20 fatty acids, sodium salts

Long and medium chain fatty acids, present as their sodium salts in TOS, are the most abundant constituents of TOS.  As their triglyceride esters (fats and oils), they are normal components of the mammalian diet.  They are hydrolysed by lipases in the gut and the fatty acids are solubilised by bile acids produced by the liver.  The fatty acid component is rapidly absorbed by the small intestine by a specific biochemical mechanism and transported in the blood as small micelle-like particles called chylomicrons.  In the tissues, they are a major source of metabolic energy; however, unsaturated fatty acids (e.g. oleic, linoleic and α-linolenic acids) are metabolised to form vital biochemical components of mammalian tissues and cell membranes. Block 1 is mostly toxicologically benign, or even positive, although, some oxidative metabolites of linoleic acid such as certain fatty acid diols produced from coronaric acid have been shown to be toxic under certain circumstances.

Medium and long chain fatty acids can be divided into the following sub-classes due to differences in details of their metabolism and their ultimate metabolic fate.

1.       C-even saturated fatty acids. These are the common fatty acids present in foods of both plant and animal origin such as stearic (C18) and palmitic (C16) acids. They are metabolised to form energy in the cell by the biochemical process known as β-oxidation.

2.       C-odd saturated fatty acids.  Example is margaric acid, which has 17 carbon atoms. Almost all animal fatty acids have an even number of carbon atoms, except for ruminant fat and milk, where they are possibly derived from microorganisms in the rumen.  However, plants produce several C-odd fatty acids, both saturated and unsaturated. Consequently, a number of these are present in TOS. They are broken down to form energy by a slightly different pathway involving propionyl-CoA rather than acetyl-CoA.

3.       Mono-unsaturated fatty acids.  An example is oleic acid, which is the second most abundant fatty acid in TOS. These are important biochemicals for cell membrane structure but are also used for energy in the same way as saturated fatty acids.

4.       Poly-unsaturated fatty acids (two and more carbon-carbon double bonds). Examples are linoleic acid (two double bonds) and linolenic acid (three double bonds), which are essential fatty acids for humans and important in the biosynthesis of arachidonic acid and eicosanoids.

5.       Most carbon-carbon double bonds in fatty acids exist in the cis-configuration. However, several trans fatty acids of plant origin occur in TOS.  An example is bovinic acid.

Fatty acids are required components in the diet of animals and are metabolised in cells to generate cellular energy in the form of adenosine triphosphate (ATP) and as such, they are the major source of metabolic energy in the western diet.  Additionally, dietary deficiencies of certain unsaturated fatty acids (essential fatty acids, which cannot be synthesised in vivo but must be obtained via the diet) such as linoleic acid (18:2 omega-6) and linolenic acid (18:3 omega-3) during human development has been shown to result in reduced levels of 20:4 omega-6 and 22:6 omega-3 fatty acids in the developing central nervous system, and this has been associated with altered learning behaviour and visual function (Innis, 1993). This is due to their function as major acyl components of cell membrane phospholipids in the central nervous system. Consequently, fatty acids may be considered as both sources of energy-rich food and essential components of the animal body.  In the normal diet, fatty acids are mainly ingested as their triglyceride esters (fats and oils) and are broken down in the small intestine (mainly the duodenum) where they are hydrolysed to mono- and di-glycerides plus free fatty acids by pancreatic lipase.  Once adsorbed, free fatty acids may either be used to make triacylglycerol, which is stored as fat as a potential energy source or broken down and catabolised to CO₂ via oxidative metabolism as an instant source of energy.  The catabolism of a saturated fatty acid such as stearic acid is rather different from that for unsaturated fatty acids (linoleic, oleic and bovinic) and will be considered separately.

Metabolism of Saturated C-even fatty acids (Applies to Palmitic (C=16) Stearic (C=18), Arachidic (C=20)

Stearic acid is desaturated to form oleic acid by the fatty acid desaturase enzyme, stearoyl-CoA 9-desaturase acting on stearoyl-CoA to the extent of 9.2% in humans (Emken, 1994). This desaturation reaction takes place in the microsomal fraction and is coupled to NAD (Kinsella, 1972).

However, stearic acid, in common with other C-even saturated fatty acids, is principally catabolised by a biochemical process known as β-oxidation. Oxidation of fatty acids takes place in the mitochondrion and happens in three stages. In the first stage, β oxidation the fatty acids undergo oxidative removal of successive two-carbon units in the form of acetyl-coenzyme A (acetyl-CoA), starting from the carboxyl end of the fatty acyl chain (Fig. 5.1.1). The 18-carbon fatty acid stearic acid undergoes eight passes through this oxidative sequence; in each pass it loses two carbons as acetyl-CoA. At the end of eight cycles the last two carbons of stearic acid (originally C-17 and C-18) are left as acetyl-CoA. The overall result is the conversion of the 18-carbon chain of stearic acid to nine two-carbon acetyl-CoA molecules. The acetyl-CoA then enters the tricarboxylic acid (Krebs) cycle and in so doing is oxidised to CO₂.

 

The 8 Successive β-Oxidation Steps of the Hydrocarbon chain for stearic acid is shown in Figure 5.1.1 attached in section 13.

During β-oxidation, four types of enzymic transformations take pace, first a dehydrogenation between the α and β carbon atoms of stearic acid, forming a trans double bond, catalysed by acyl-CoA dehydrogenase to give trans-Δ2-enoylCoA and utilising FAD as the hydrogen acceptor.  The reduced FAD then donates its electrons to the electron carrier electron-transferring flavoprotein, which is an integral protein of the inner mitochondrial membrane. These are then transferred via the electron transport proteins and redox compounds of the inner mitochondrial membrane (complex II→ ubiqinone → complex III → cytochrome c → complex IV → cytochrome oxidase → O₂).  This results in 148 mol of ATP synthesised per mol of stearic acid, showing why fatty acids are such good sources of metabolic energy and compares with about 30 mol ATP per mol glucose.  Hence, fats produce about 3-fold more metabolic energy than carbohydrates on a mass basis.  Consequently, in animals, fat in adipose tissue is the major long-term storage medium for energy and a high proportion of ingested fatty acids are metabolised to form fat.

Unlike other saturated fatty acids, dietary stearic acid does not appear to raise plasma cholesterol levels. The reason for this unclear, although it appears that it be related to inherent differences in the metabolism of the fatty acids. The comparative metabolism of palmitic, stearic and oleic acids by cultured hamster hepatocytes was investigated by Bruce and Salter (1996). In this study, stearic acid was taken up more slowly and was poorly incorporated into both cellular and secreted triacylglycerol than the other two fatty acids. Despite this, stearic acid stimulated the synthesis and secretion of triacylglycerol to the same extent as the other fatty acids. Incorporation into cellular phospholipid was lower for oleic acid than for palmitic acid and stearic acid. Desaturation of stearic acid, to oleic acid, was found to be greater than that of palmitic acid to the corresponding mono-unsaturated fatty acid. Oleic acid produced from stearic acid was incorporated into both triacylglycerides and phospholipid, representing 13% and 6% respectively of the total after a 4 h incubation. Significant proportions of all the fatty acids were oxidized, primarily to form ketone bodies, but by 8 h, more oleic acid had been oxidized compared with palmitic acid and stearic acid.

Metabolism of Saturated C-odd Fatty Acids (Applies to Margaric Acid (C17 and 21:0 Fatty Acid))

The major difference in metabolism of odd-numbered fatty acids is that when they are oxidized via β-oxidation, they produce propionyl CoA and acetyl CoA in the final round of degradation rather than two molecules of acetyl CoA. The activated 3 carbon unit in propionyl CoA, once converted into succinyl CoA, enters the citric acid cycle. The biotransformation of propionyl CoA to succinyl CoA is effected by the three enzymes propionyl-CoA carboxylase, methylmalonyl-CoA epimerase and methylmalonyl-CoA mutase, which are required to complete the metabolism of C-odd fatty acids (Wongkittichote et al, 2017).  

Metabolism of Mono-unsaturated Fatty Acids (Applies to Oleic (C=18) Palmitoleic (C=16), 20:1 Fatty acid, 18:1 Fatty acid (11)

Oleic acid is a very common unsaturated fatty acid and as its triglyceride occurs in appreciable quantities in most of the common cooking oils of commerce. Olive oil contains particularly high amounts (55% to 83%). Oleic acid is metabolised (oxidised) in the mitochondria by an identical metabolic pathway to linoleic acid. It is the second most abundant constituent of Block 1 in TOS.  

In general, studies on the oxidation (catabolism) of fatty acids have shown that long-chain fatty acids are oxidised more slowly, and unsaturated fatty acids are oxidised more rapidly than saturated fatty acids. Measurement of fatty acid oxidation in rats with a series of fatty acids showed that oxidation of the saturated fatty acids decreases with increasing carbon length (laurate> myristate > palmitate > stearate) (Leyton et al. 1987)). For unsaturated fatty acids, 24-h oxidation was in the following order: linolenate >oleate > linoleate > arachidonate. Through 7 h, the oxidation of oleate was greater than that of linolenate. Thus, the medium chain fatty acids (8–14 carbons) were oxidized the most rapidly, with linolenate and oleate oxidation occurring nearly as rapidly. This suggests that unsaturated fatty acids are more directed towards the mitochondria for oxidation than saturated fatty acids, which in turn are stored preferentially as triacylglycerol (fat). Hence, oleic acid is more rapidly processed through mitochondrial β-oxidation than stearic acid and less likely to be sequestered as fat.  See also DeLany et al. (2000).

Oleic acid is considered an important dietary fatty acid and its incorporation into lipid membranes is considered vital with respect to the membrane structure, fluidity and function and the action of the important G-protein mediated protein receptors embedded within the membrane and having seven transmembrane loops (Lopez et al, 2014). The ligands that bind and activate these receptors include light-sensitive compounds (retinaldehyde), odours, pheromones, hormones, and neurotransmitters. The ligands vary in size from small molecules to peptides to large proteins. G protein-coupled receptors are involved in many diseases and are also the targets of a high proportion of modern medicinal drugs (Hauser et al, 2017).

Metabolism of Poly-unsaturated Fatty Acids (Applies to Linoleic acid, α-Linolenic acid, 20:3 fatty acid, Palmitoleic acid)

Linoleic (an ω-6 dienoic acid) is the most abundant fatty acid in TOS and it is one of the two truly essential fatty acids in humans (the other is the ω-3 trienoic acid, α-linolenic acid). It is essential in the human diet, probably because it is the substrate for the synthesis of longer-chain, more unsaturated ω-3 fatty acids, as is α-linolenic acid.  The initial metabolic step is via the action of acyl-CoA-desaturase to yield γ-linolenic acid (Fig. 5.1.2 attached to section 13). Humans cannot synthesis linoleic acid or α-linolenic acid because they lack the desaturase enzyme required to make them from oleic acid.  Linoleic is modified metabolically to the following groups of vital cell components: eicosanoids (affecting inflammation and many other cellular functions), endocannabinoids (affecting mood, behaviour and inflammation), lipoxins which are a group of eicosanoid derivatives formed via the lipoxygenase pathway and several other important cellular components.

The metabolism of linoleic acid in humans to form arachidonic acid (C20:4n-6), which is a major precursor of eicosanoids has been investigated by Vermunt (Vermunt et al. 2001). Linoleic methyl ester was administered orally in an olive oil solution.  The 13C-enriched metabolites were identified by GC/MS. Both blood samples and breath (13CO₂) were monitored for 13C-containing metabolites.  The highest concentration of fatty acid in plasma was the base material linoleic acid (1.12% of the applied dose), which was reached after16.2 hr on average. Appearance of γ-linolenic acid (13C18:3n-6) in the plasma occurred almost at the same time as that of linoleic acid (13C18:2n-6), showing a high level of Δ5-6 desaturase.  The peak of arachidonic acid (C20:4n-6) was reached about 160 hr after administration. See Fig. 5.1.2 for the metabolic pathway from linoleic acid to arachidonic acid.

Peaks in ¹³C enrichment in breath were reached after 3 ± 5 h and ranged from 7 ± 12%.  After 6 h, total 13C recovery ranged between 11.9 and 17.4% while after 12 h total recovery of the tracer was increased to 16.8 ± 25.1%.

Consequently, linoleic acid metabolism comprises both a rapid catabolic mechanism via β-oxidation and the Krebs cycle to yield CO₂ (major route, see above) and a minor, but very important route consisting of Δ5-6 desaturase steps alternating with 2-carbon chain elongation steps to yield arachidonic acid.  

The human metabolism of α-linolenic acid has been reviewed by Burdge (Burdge, 2006), who suggests that the reason that it is an essential fatty acid is due primarily to it being a substrate for the synthesis of the long-chain, more unsaturated poly-unsaturated fatty acids (PUFA), eicosapentaenoic acid (20:5n-3) and docosahexaenoic acid (22:6n-3) (Fig. 5.1.3 attached to section 13). As an important building-block for other biochemicals, the amount used for metabolic fuel via β-oxidation is less for α-linolenic acid. The proportion of [13C] α-linolenic acid that is consumed in β-oxidation was 33% on average of the administered dose in men (Burdge et al. 2002, Bretillon et al, 2003, Burdge et al, 2003, DeLany et al, 2000, Vermunt et al. 2000) and 22% in women (Burdge and Wootton, 2002).

The metabolism of linoleic acid to arachidonic acid is shown in Figure 5.1.2.

The metabolism of Linolenic Acid to Longer Chain PUFAs is shown in Figure 5.1.3.

β-Oxidation of Unsaturated Fatty Acids (Applies all C-even cis Unsaturated Fatty Acids).

A very high proportion of unsaturated fatty acids (linoleic and oleic) are metabolised in the mitochondrion to provide energy; however, the ratio is less than for saturated acids such as stearic or palmitic acids (Bakke et al, 2012, McCloy et al, 2007), as would be expected due to the alternative routes of unsaturated fatty acid metabolism.

There is a complicating factor in the β-oxidation of unsaturated fatty acids and that  the carbon-carbon bonds are in the cis configuration and cannot be acted upon by enoyl-CoA hydratase, the enzyme catalysing the addition of water to the trans double bond of the Δ2-enoyl-CoA generated during β oxidation. However, by the action of two auxiliary enzymes, the fatty acid oxidation sequence can also break down unsaturated fatty acids. These two enzymes are enoyl-CoA isomerase, which makes the cis double bond trans and converted by enoyl-CoA hydratase into the corresponding L-β-hydroxyacyl-CoA (trans-Δ2dodecenoyl-CoA) (see Fig. 5.1.4 attached to section 13).

The β-Oxidation of an Unsaturated Fatty Acid (Applies to all Unsaturated Fatty Acids) is shown in Figure 5.1.4

Peroxidation and Oxidation of Unsaturated Fatty Acids

Additionally, all unsaturated fatty acids are subject to oxidation via a free radical mechanism that initially gives hydroperoxides, however, this is generally only of toxicological importance when the polyunsaturated fatty acids are present in cellular membranes.  In summary, cell injury activates lipoxygenases enzymes, which generate lipid hydroperoxides and in the presence of Fe, free radicals, fatty acid free radicals are produced. Radicals will attack any activated CH2-group on polyunsaturated fatty acids. Since linoleic acid is the most abundant polyunsaturated fatty acid in mammals, its lipid peroxidation products dominate. Lipid peroxides are reduced in biological systems to the corresponding hydroxy acids (LOHs). LOHs derived from linoleic acid, hydroxyoctadecadienoic acids (HODEs), are the main marker of lipid peroxidation. In the case of linoleic acid, the main peroxidation product is 8-hydroxy-9Z,12Z)-octadeca-9,12-dienoic acid (Fig. 5.1.5 metabolite 1). HODEs are of high physiological relevance and lipid peroxidation is implicated in several disease states mediated by oxidative stress such as a number of cardiovascular, infectious and genetic diseases. For a review on lipid peroxidation see Spiteller (1998).

In addition to peroxidation, linoleic acid is subject to oxidation of its carbon-carbon double bonds to form oxiranes (epoxides). This is generally catalysed by several specific cytochromes P45.  A significant metabolite derived from linoleic acid is coronaric acid (9,10-epoxy-12Z-octadecenoic acid, Fig. 5.1.5 metabolite 2 attached in section 13), which is a significant urinary metabolite (Konkel and Schunck , 2011). This metabolite is also produced non-enzymatically by autoxidation if linoleic acid  is exposed to oxygen and/or UV radiation. Further metabolism leads to the formation of two diastereomeric dihydroxy fatty acids in mammalian tissue 12S,13R-dihydroxy-9(Z)-octadecaenoic and 12R,13S-dihydroxy-9(Z)-octadecaenoic acids by a soluble epoxide hydrolase within minutes of its formation (Greene et al, 2000).  The metabolism of coronaric acid to these two dihydoxy fatty acids, which are collectively termed isoleukotoxin diols (Fig. 5.1.5 metabolite 4) appears to be critical for the toxicity of coronaric acid, i.e. the diols are the toxic metabolites of the less toxic coronaric acid.  It has also been reported that the isomeric form 12,13-epoxy-9Z-octadecenoic acid is also formed on oxidation of linoleic acid (Fig. 5.1.5 metabolite 3) and on hydrolysis by epoxide hydrolase yields the isomeric diols 9S,10R-dihydroxy-12(Z)-octadecaenoic and 9R,10S-dihydroxy-12(Z)-octadecaenoic acids, which are known as leucotoxin (Fig. 5.1.4 metabolite 5) (Greene and Hammock, 1999b). Cytotoxicity is associated with mitochondrial dysfunction with respiration decreasing 54% prior to the onset of cell death. Secondary to the mitochondrial toxicity, the diols completely inhibited active Na+ transport. (Moran et al., 1997, Greene and Hammock, 1999a).

The peroxidation and oxidation of linoleic acid is shown in Figure 5.1.5.

Unsaturated Fatty Acids Containing trans Carbon-Carbon Double Bonds (Applies to Bovinic acid and 18:2 fatty acid (t,t))

Bovinic acid ((9Z,11E)-octadeca-9,11-dienoic acid) also known as rumenic acid is a conjugated linoleic acid found in the fat of ruminants and in dairy products. It is an omega-7 trans-fat and is found mostly in milk and milk products but can also be synthesized in mammalian tissues from trans-vaccenic acid (C18:1 t11) through the action of delta-9 desaturase (Schneider et al, 2017). Bovinic acid is attributed with several health promoting effects, including anti-cancer and anti-atherosclerotic activities.  The degradative metabolism would be expected to be similar to that of linoleic acid (see above), although definitive information could not be found. There is evidence that these naturally occurring trans fatty acids do not have the deleterious effects that synthetic trans fats produced by the industrial partial hydrogenation of vegetable oils have: e.g., a study by the US Department of Agriculture showed that vaccenic acid raises both high density lipoprotein (HDL) and low density lipoprotein (LDL) cholesterol in humans, whereas industrial trans fats only raise LDL with no beneficial effect on HDL (Baer, 2010, Gebauer et al, 2010).

Block 2: Rosin acids and their sodium salts

Rosin acids, such as abietic acid, contribute the second most abundant component in TOS.   They are present as their sodium salts in TOS. They are common components in all coniferous trees. Some of the components are double bond isomers and can inter-convert in vivo and in the environment. They are rapidly conjugated and excreted in the urine.

Abietic acid (1R,4aR,4bR,10aR)-1,4a-dimethyl-7-propan-2-yl-2,3,4,4b,5,6,10,

10a-octahydrophenanthrene-1-carboxylic acid) is the principal diterpenoid rosin acid produced in the oleoresin of many coniferous (softwood) trees of the genera Pinus (pines), Picea (spruces) and Abies (firs); consequently, it would be expected to be present in TOS derived from wood pulping. Other rosin acids commonly produced include pimaric acid, palustric acid, dehydroabietic acid, neoabietic acid and isopimaric acid. Several of these compounds are double bond isomers (palustric, abietic and neoabietic) and may be inter-converted in the environment via acid catalysed isomerisation (Morales et al., 1992). Abietic and the other rosin acids are all hydrophobic compounds with solubilities of less than 6 mg/L in water at neutral pH (Nyren and Black, 1958).

Metabolism of rosin acids

Data on the metabolism of abietic acid have been taken from the MAK Collection (Wiley On-line Library, 2103).  This document summarises the chemical’s physical properties, uses, and metabolism, effects in humans and animals, amongst other effects and chemical properties.

Interconversion of abietic and Other rosin acids

Other rosin acids found in TOS are pimaric acid, palustric acid, dehydroabietic acid, neoabietic acid and isopimaric acid. Several of these compounds are double bond isomers (palustric, abietic and neoabietic) and may be inter-converted in the environment via acid catalysed isomerisation (Morales et al., 1992). Abietic acid is in an equilibrium state with levopimaric acid, neoabietic acid and palustric acid, which shifts to the side of abietic acid from 100°C; from 250°C irreversible conversion to mostly dehydro-, but also dihydro- and tetrahydroabietic acid (disproportionation) takes place (Song et al. 1985).  Consequently, the presence of levopimaric acid, neoabietic acid, palustric acid and dehydroabietic acid, all of which are found in TOS, would be expected (Fig. 5.1.6 attached in section 13). Consequently, commercially available abietic acid has a purity of 85% to 90% (Karlberg 1989) or 74% with 13.6% isopimaric, acid and 7.5% dehydroabietic acid (Karlberg et al. 1985).

The interconversion of rosin acids is shown in Figure 5.1.6

Oxidation of rosin acids

Abietic acid and other rosin acids are also subject to air-oxidation and abietic acid and dehydroabietic acid can contain oxidation products, e.g. 13,14-α-epoxides, 13,14-β-epoxides, 7-oxo- or 15-hydroperoxy compounds (see Figure 5.1.7 attached; Karlberg and Gäfvert 1996; Karlberg et al. 1988 a, b), although none were detected in the samples of TOS analysed.

The air oxidation of abietic acid is shown in Figure 5.1.7

Abietic acid

Six male rabbits were given 2000 mg abietic acid sodium salt (about 666 mg abietic acid sodium salt/kg body weight at 3 kg body weight) via gavage as suspension in an aqueous Tween-80 solution. Abietic acid was oxidised on the isopropyl group at C-17, probably via a primary alcohol as intermediate (metabolite 1 in Fig. 5.1.6) to form an acid (metabolite 2, Fig. 5.1.6), which was 27% of the acid ether extract of the urine. Only urine metabolites were investigated (The MAK‐Collection for Occupational Health and Safety, 2013, Asakawa et al. 1986). These metabolites are shown in Fig. 5.1.8.

The degradative metabolic pathway of abietic acid in rabbits is shown in Figure 5.1.8

It is suggested that the uptake, toxicokinetics and metabolism of neoabietic acid and palustric acid may parallel those of abietic acid, as these compounds are inter-convertible (see above).

Dehydroabietic acid

The uptake and metabolism of (+)-dehydroabietic acid in rabbits was investigated by Matsumoto et al. (1990).   Six male Japanese White Strain rabbits were orally administered a total of 12 g of sodium dehydroabietate (purity and/or analysis not stated) as a 100 mg/mL suspension in 0.02% sodium sorbitate containing Tween 20 surfactant.  Urine was collected over a period of 3 days in a pH 4 aqueous solution.  The urine solution was treated with a mixture of β-glucuronidase/arylsulfatase to convert the phase II urinary conjugates (glucuronides and sulfates) to their aglycones. These were converted to their methyl esters using diazomethane and separated by flash chromatography on a silica gel column using mixtures of hexane, benzene, and ether as mobile phase.  The total yield of urinary metabolites was 40% of the administered dose.  The structures of the 10 metabolites isolated were determined by 1H NMR, IR and UV spectroscopy EI mass spectroscopy and chemical synthesis of derivatives. The absolute stereochemical configuration for 7 out of the 10 metabolites were determined. The 10 metabolites are summarised in the table below and their numbering is the same as for Fig. 5.1.9.

Table 5.1.2: Urinary Metabolites of dehydroabietic acid in rabbits

Metabolite No.	Chemical Name	Recovery Yield (%)
1	8,11,13,15-abietatetraen-18-oic acid	2.28
2	15-hydroxy-8,11,13-abietatrien-18-oic acid	2.42
3	2β,15-dihydroxy-8,11,13-abietatrien-18-oic acid	1.49
4	2α,15-dihydroxy-8,11,13-abietatrien-18-oic acid	2.56
5	(15R)-15,16-dihydroxy-8,11,13-abietatrien-18-oic acid	0.62
6	2α-hydroxy-8,11,13,15-abietatetraen-18-oic acid	3.23
7	(15S)-8,11,13-abietatrien-16,18-dioic acid	0.45
8	16-hydroxy-8,11,13-abietatrien-18-oic acid	0.20
9	(15S)-2β,16-dihydroxy-8,11,13-abietatrien-18-oic acid	0.54
10	(15S)-2α-dihydroxy-8,11,13-abietatrien-18-oic acid	0.27

The Urinary Metabolites of Dehydroabietic Acid in Rabbits is shown in Table 5.1.9

Orally-administered dehydroabietic acid was rapidly and efficiently (40% yield) metabolised to yield 10 oxygenated metabolites as their conjugates. The primary route of hydroxylation was by transformation of the isopropyl group and secondarily by ring hydroxylation in the 2-position (see metabolic pathway below).  The authors suggest that that the oxidation of dehydroabietic acid in rabbits occurs first at C-15 and C-16 to give the metabolites 2 and 8. The evidence indicates that the dihydroxyisopropyl metabolite 5 is formed directly from dehydroabietic acid; although alternative routes from 2 and 8 are also possible. It is suggested that the 13-isopropenyl derivatives 1 and 6 are artefacts and are formed from the dehydration of the tertiary alcohols 2 and 4 respectively under the acidic conditions (pH 4).

The earlier paper by Asakawa studied the metabolism of both (+)-dehydroabietic acid and (-)-abietic acid, administered as their sodium salts (Asakawa et al, 1986). The metabolites of dehydroabietic acid were isolated were 1, 2 and 8 of those identified in the later Matsumoto et al, (1990) paper.  In addition, the 7-oxo derivative of dehydroabietic acid was also identified (shown as metabolite 11 in Fig. 5.1.9) as a trace metabolite. This metabolite was not identified in the later paper (Matsumoto et al. 1990).

The mammalian toxicokinetics of dehydroabietic acid seem to be highly dependent on the species tested, as when rats were administered [3H]-dehydroabietic acid only 5-7% of the radioactivity was excreted in the urine, with the bulk (80-90%) being passed in the faeces. However, the suggestion is that the bulk of the material passed in faeces was via alimentary canal uptake and re-excretion via the liver and bile (see below).

After oral administration of 2 or 300 mg [3H]-dehydroabietic acid/kg body weight to rats, 80% to 90% of the administered radioactivity was excreted with the faeces and 5% to 7% with the urine within 2 days. Of the residual radioactivity in the body, the major part was in the gastrointestinal tract and in the carcass. After administration of 150 mg [3H]-dehydroabietic acid/kg body weight, up to 40% of the recovered radioactivity was found in the gall bladder in rats with bile duct fistula. From this, the authors concluded that up to 40% of the administered [3H]-dehydroabietic acid was absorbed via the gastrointestinal tract. After administration of 150 mg [3H]-dehydroabietic acid/kg body weight, assays taken at 2, 4 and 6 hours thereafter revealed the radioactivity distributed in the tissues. (The MAK‐Collection for Occupational Health and Safety, 2013, HSE, 1998).

Pimaric acid and isopimaric acid

Isopimaric and tetrahydroabietic acid were also absorbed after oral administration [to rats] and mostly excreted with the faeces. (The MAK‐Collection for Occupational Health and Safety, 2013, HSE, 1998).

No experimental data could be found on the metabolism of pimaric acid in mammals.

Block 4: Terpenes

α-Pinene

The chirality of α-pinene varies with the species of pine tree. In European pines, it is of the (-) 1S,5S configuration and North American pines it is (+) 1R,5R. Metabolic studies often do not state the isomer composition of the α-pinene used; however, this can influence the nature of the metabolites formed (e.g. Ishida et al., 1981). The metabolic transformations of α-pinene have been determined by White et al (1979) in rat liver microsomes, Ishida et al. (1981) in rabbits, and in humans by Eriksson and Levin, (1990, 1996) and Schmidt et al. (2013). Alpha-pinene is an inducer of cytochrome P450 enzymes in mammals and its principal route of phase I metabolic transformation is the production of various hydroxylated terpenols shown in the metabolic pathway scheme below, catalysed by cytochromes P450.

 

An in vitro study using rat liver microsomes identified α-pinene oxide [1] as a principal metabolite (White et al., 1979). Epoxides such as [1] are frequent intermediates in the hydroxylation of alkenes. In a series of studies in which urine from sawmill workers exposed to α-pinene, β-pinene and d-3-carene via their work was collected and hydrolysed with beta-glucuronidase, and the resultant aglycones analysed by GC/EI-MS, the principal metabolites were identified exclusively as those derived from α-pinene; no metabolites of β-pinene and d-3-carene were identified. Note, although environmental monitoring of α-pinene, β-pinene and d-3-carene was carried out, the concentrations of these materials to which the workers were exposed in the experiment were not reported. The metabolites identified were cis- [2] and trans-verbenol [3]. The recoveries of the verbenols from hydrolysed urine were in the range of 85 to 94%. These metabolites were not detectable in urine that was not treated with glucuronidase, indicating that the two verbenols are excreted in urine as the glucuronides (Eriksson and Levin, 1990).  In a follow-up study the same authors analysed the urine of similarly exposed workers using EI and CI MS post enzymatic hydrolysis and GC separation following TMS derivatisation (Eriksson and Levin, 1996). In addition to the cis-and trans-verbenol identified in the earlier study, they showed evidence for the production of a transdiol derivative of trans-verbenol [metabolite 4] and its alcohol-aldehyde oxidation product [metabolite 5]. The diol derived from cis-verbenol [metabolite 6] was also identified.

The metabolism of α-pinene in laboratory rabbits was reported by Ishida (Ishida et al., 1981). Rabbits were orally dosed with (+)- (-)- and (±)-a-pinene and the urinary metabolites subjected to glucuronidase and arylsulfatase digestion. Metabolites were purified by silica gel chromatography, GC and TLC. Pure metabolites were identified by EI MS and 1H-NMR. Myrtenol [7] and its oxidation product myrtenic acid [8] were identified. The yields of myrtenol [7] were highly dependent of the nature of the chirality of the α-pinene used: (+) 14.9% (-) 0.9% (±) 9.3%. Myrtenol [7] has also been identified in the urine of human volunteers orally exposed to a cough medication containing monoterpenes, including α-pinene, as the active medicinal ingredients (Meesters et al., 2008).

The metabolites of α-pinene isolated from the urine of humans who had not been occupationally exposed to monoterpenes using a highly sensitive GC MS/MS method following TMS derivatisation were (1S,2S,5S)-cis-verbenol, (1R,2S,5R)-trans-verbenol and (1S,5R)-(+)-myrtenol [7] (Schmidt et al., 2013). Borneol, bornylacetate and myrtenol [7] have been identified as metabolites from α-pinene in human urine after acute poisoning with pine oil (Koppel et al.,1981). The biosynthesis of borneol structures from α-pinene seems unlikely considering the uncertainty in identification of borneol and lack of mechanistic explanation for their biosynthesis. Therefore, these structures are not shown in Figure 5.1.10. A composite pathway for α-pinene metabolism in mammals is shown in Figure 5.1.10.

β-Pinene

In contrast to α-pinene, the isomer of β-pinene in different conifer species is always (-)- β-pinene. Ishida et al. (1981) considered the principal route of (-)-β-pinene metabolism in rabbits to be via the allylic oxidation of the alkene moiety to form an epoxide intermediate [metabolite 1, Figure 5.1.11 attached in section 13]. This was further ring-opened and hydrolysed to yield (-)- α-terpineol [2] and (-)-1-p-menthene 7,8 diol [3] in 39% and 30% yield respectively, the magnitude of the yield implying that this is the major catabolic route.  (+)-Trans-pinocarveol [4] and (-)-trans-10-pinanol [5] formed by ring and alkene hydroxylation respectively of β-pinene were relatively minor metabolites. In the brushtail possum (Trichosurus vulpecula) myrtenic acid was identified as a non-conjugated metabolite, which was excreted in the urine after oral administration of β-pinene (Southwell et al.,1980). A composite pathway for β-pinene metabolism in mammals is shown in Figure 5.1.11.

Block 5: Sesquiterpenes

 γ-Muurolene

Muurolene is a sesquiterpene that is used in the perfumery industry.  No experimental data on its in vivo uptake and metabolism could be located, however, an in silico study was used to predict the likely metabolites in mammals (Silva et al, 2017). MetaPrint2D-React online tool was used to predict the metabolic pathways of γ-muurolene with simulations in phase I and II of potential human, dog and rat bio-transformations. The predictive oxygenated metabolites were via oxidation of the 15-methyl group to give sequentially the aldehyde (1), the primary alcohol (2) and finally the carboxylic acid (3). It was predicted that the number 3 ring carbon would be metabolised to the secondary alcohol (4), which would give rise to the ketone (5).  Finally, it was predicted that diene (6) would be a metabolite in addition to the reduction of the 4,5 double bond to give (7, cadalane). γ-Muurolene and the metabolites were not predicted as toxic regarding the hepatotoxicity and hERG inhibition. However, a predicted risk of bacterial mutagenicity was detected for metabolite 7, presumably due to possible epoxide formation.

The in-silico predicted metabolites of γ-muurolene is shown in Figure 5.1.12

γ-Cadinene

γ-Cadinene is a sesquiterpene that is common in the essential oils of many plants, including conifers. No experimental data could be found on the mammalian toxicokinetics and metabolism of γ-cadinene.

Block 6: Abietenes and Labdanes

Abietadiene

Abietadiene is a biosynthetic intermediate of abietic acid in conifers (MetaCyc Pathway: abietic acid biosynthesis, 2009).  It is produced by the isoprenoid biosynthesis pathway starting from (E,E,E)-geranylgeranyl pyrophosphate (1) to yield the unstable tertiary labdane alcohol 13-hydroxy-8(14)-abietene (2) by the action of abietadiene synthase (Keeling et al, 2011, Zou et al. 2012). Metabolite 2 is then dehydrated to yield abietadiene (3) as well as lesser amounts of its double bond isomers (-)-levopimaradiene, (-)-neoabietadiene and (-)-palustradiene (structures not shown). Abietadiene is then sequentially oxidised at the 18-position to afford abietadienol (4), abietadienal (5) and abietic acid (6) via the action of three NAD or NADP-linked dehydrogenases (MetaCyc Pathway: abietic acid biosynthesis, 2009). No direct experimental data could be found on the mammalian toxicokinetics and metabolism of abietadiene, but it is likely to follow a similar route to abietic acid, although the level of uptake is likely to be much less in mammals due to its high log K_ow.

The biosynthesis of abietadiene and its conversion to abietic acid in conifers is shown in Figure 5.1.13

Labdane

No experimental data could be found on the mammalian metabolism of labdane, although the level of oral uptake is likely to be low in mammals due to its high log K_ow.

Thunbergene

(+)-Thunbergene ((+)-Cembrene), a diterpene hydrocarbon, occurs in a number of plant species including Pinus koraiensis (Korean pine) and marine corals. It has been previously identified as a constituent of effluent from the pulping process (Koistinen et al, 1998). No experimental data could be found on the mammalian metabolism of thunbergene.

Block 7: C30 branched polyalkenes

 

Squalene

Squalene, an isoprenoid compound, is an intermediate metabolite in the synthesis of cholesterol. Its biosynthesis is via the isoprenoid pyrophosphate pathway, whereby two molecules of farnesyl pyrophosphate condense to form squalene, which is catalysed by squalene synthase and reduction by NADPH.  It is present in appreciable amounts in olive oil (36-708 mg/100g) (Gutfinger and Letan, 1974)

The level of oral uptake of squalene would be expected to be low in mammals due to its high log K_ow, however, this is not so, and dietary squalene is absorbed efficiently by the small intestine. In humans, between 60 and 85% of dietary squalene is absorbed from an oral dose (Bargossi et al, 1994, Strandberg et al, 1990, Miettinen and Vanhanen, 1994, Gylling and Miettinen, 1994). It is transported in the blood plasma, generally in association with very low-density lipoproteins (LDL) and is distributed widely in human tissues (Strandberg et al, 1990), with the greatest concentration in the skin, where it is one of the major components of skin surface lipids. For an overview see Kelly (1999).

In humans, feeding squalene had no consistent effect on absorption efficiency of cholesterol, but significantly increased the faecal excretions of cholesterol and its nonpolar derivatives coprostanol, epicoprostanol, and coprostanone and bile acids, indicating an increase of cholesterol synthesis by about 50% (Strandberg et al, 1990). These results imply that a substantial amount of dietary squalene is absorbed and converted to cholesterol, but this squalene-induced increase in synthesis was not associated with consistent increases of serum cholesterol levels. The authors suggest that increased levels of esterified methyl sterols measured in the blood serum may reflect stimulated tissue acyl CoA: cholesterol acyltransferase activity during squalene feeding as these sterols are not esterified in serum.

The absorption and metabolic fate of dietary squalene was investigated on the rat by administering a single oral dose of ³H-squalene and ¹⁴C-cholesterol (Tilvis and Miettinen, 1983). Experiments on rats with a cannulated thoracic duct revealed that ³H-squalene was, like ¹⁴C-cholesterol, absorbed through the lymphatic vessels and that about 20% of absorbed ³H-squalene was cyclized to sterols during the movement through the intestinal wall. Faeces contained ³H-sterols, indicating that newly synthesized mucosal sterols had been secreted into the gut lumen. In intact animals, ³H-squalene appeared in the circulation more rapidly than ¹⁴C-cholesterol and did not persist to any significant extent in the squalene-rich adipose and muscle tissues. The increase in dietary squalene load (8-48 mg) decreased the absorption percentage of 3H-squalene (45-26%) but did not affect the absorption of ¹⁴C-cholesterol (47%). Measurement of faecal steroids revealed that initially absorbed ³H-squalene was eliminated to a significantly higher extent than ¹⁴C-cholesterol as faecal bile acids (34% vs 11%). The experiments showed that the rat intestine has a marked capacity for absorbing dietary squalene and that the absorbed squalene is preferentially converted into bile acids in the liver.

Squalene is not very susceptible to peroxidation and appears to function in the skin as a quencher of singlet oxygen, protecting human skin surface from lipid peroxidation due to exposure to UV and other sources of ionizing radiation. Squalene may also act as a "sink" for highly lipophilic xenobiotics. Since it is a nonpolar substance, it has a higher affinity for non-ionised drugs. In animals, supplementation of the diet with squalene can reduce cholesterol and triglyceride levels. In humans, squalene has the potential to be a useful addition to potentiate the effects of some cholesterol-lowering drugs.

Squalene is a well-known dietary component and biochemical intermediate and its metabolism to give cholesterol, bile acids and other steroids is well documented in biochemical textbooks.

Block 8: 3,5-dimethoxystilbene

3,5-Dimethoxystilbene is one of several related stilbene compounds that have been found in conifers, which explains its presence in TOS.  It weakly stimulates the synthesis of the detoxifying enzymes cytochromes P450.  It is structurally related to the antioxidant compounds resveratrol and pterostilbene, which have been extensively investigated from a biochemical aspect. From a chemical structure point of view, it is likely that 3,5-dimethoxstilbene is hydroxylated to pterostilbene, which is then rapidly conjugated as its glucuronide and excreted in mammals.

3,5-Dimethoxystilbene is one of several related stilbene compounds that have been found in conifers, especially Pinus spp. For example, 3,5-dimethoxystilbene, together with 3,5-dihydroxy- and 3-hydroxy, 5-methoxystilbene have all be identified in the wood of Pinus sylvestris and all three have been implicated in inducing cytochrome P450 activity in vitro using ethoxyresorufin-O-deethylase (EROD) activity in a rat hepatoma cell line and vitellogenin induction in fish (Parrott et al, 2011). Although 3,5-dimethoxystilbene had the highest activity of these three stilbenes, the activities of all three were weak compared with that of the rosin acid metabolite retene, which was used as a positive control.

No information could be found on the metabolism of 3,5-dimethoxystilbene; however, the compound is structurally related to the well-known anti-oxidants pterostilbene (3,5-dimethoxy-4’-stilbenol) and resveratrol (3,5,4′-trihydroxystilbene), although it is less polar.  The oral bioavailability of resveratrol is poor (Assensi et al, 2002), probably due in part to its low water solubility, and is limited by poor absorption and first-pass metabolism: only low plasma concentrations of resveratrol were seen following oral administration, and metabolism to 4’-glucuronide and 4’-sulfate conjugates was rapid via phase II metabolism in rats. When administered orally, pterostilbene demonstrated greater bioavailability and higher total plasma levels of both the parent compound and metabolites than did resveratrol. Resveratrol was not detectable following dosing with pterostilbene. Consequently, pterostilbene does not appear to act as a prodrug for resveratrol via demethoxylation (Kapetanovic et al, 2011).  

The uptake of 3,5-dimethoxystilbene is likely to parallel resveratrol although it is more hydrophobic. In the intestine, resveratrol is absorbed by passive diffusion or by forming complexes with membrane transporters, such as integrins. Once in the bloodstream, resveratrol is found in three different forms: glucuronide, sulfate or free. The free form can be bound to albumin and lipoproteins such as LDL. These complexes can be dissociated at cellular membranes that have receptors for albumin and LDL, leaving the resveratrol free and allowing it to enter cells (Delmas et al, 2011).  In humans, the poor bioavailability of resveratrol seems more to do with rapid phase II metabolism and conjugate formation than low uptake, as after a single oral dose of 25 mg was administered it was difficult to detect the non-metabolized resveratrol in circulating plasma. However, estimates of the plasma concentrations of resveratrol plus total metabolites were considerably higher: around 400–500 ng/mL (≈2 µM), indicating a very low oral bioavailability of free resveratrol, but a significant one of its metabolites (Walle et al, 2011).

It is clear from the literature that pterostilbene is rapidly metabolised in humans and rats to its 4’-O glycoside and sulfate conjugates and that evidence of 3-O demethylation could not be found.  The in vitro xenobiotic metabolism of trans-stilbene by rat and human liver microsomes and human recombinant cytochromes P450 1A1 and 1A2 has been investigated by Sanoh et al (2002), who showed that trans-stilbene was oxidised to 4-hydroxy-trans-stilbene and 4,4’-dihydroxy-trans-stilbene. Consequently, is proposed that the catabolic route of 3,5-dihydroxystilbene catabolism in rats and humans is via 4’-hydroxylation to pterostilbene (metabolite 1 in the scheme below), followed by formation of the O-glucuronide and sulfate conjugates (metabolite 2).

The proposed route of degradative metabolism of 3,5-dimethoxystilbene in rats and humans is shown in Figure 5.1.14.

Block 9: Rosin alcohol and aldehydes isomers

The constituents in this block, pimaral, abietol and agathadiol have all been reported as occurring in coniferous trees and it is likely that all are either on the synthesis or degradative pathways for the rosin acids,  However, no information on their degradative metabolism in mammals could be found, although they are likely to behave similarly to the rosin acids.

Pimaral

The sesquiterpene (+)-pimaral has been extracted from the bark of Pinus sylvestris and has been identified as a component in the sex attractant pheromone of the cerambycid beetle Monochamus galloprovincialis, a forestry pest.  This compound is likely to be on the metabolic pathway for the biosynthesis or degradation of abietic acid, but no information on the uptake, toxicokinetics or degradative metabolism could be found.

Abietol

Abietol is a biosynthetic intermediate of abietic acid (metabolite 4 in Fig. 5.1.7 above) so its degradative metabolism is likely to follow that for abietic acid in mammals.  

Agathadiol

No information could be found on the uptake, toxicokinetics and degradative metabolism of agathadiol in mammals.

Block 10: C20-C35 alcohols and terpene alcohols

Long chain alcohols are metabolised oxidatively to the corresponding fatty acids. When 1-[14C]hexadecanol was administered to cultured human fibroblast cells it was rapidly oxidised to 1-[14C]palmitic acid (Rizzio et al. 1987). This reaction is catalysed by the enzyme fatty alcohol NAD+ oxidoreductase. In vivo, the reaction is reversible and radiolabelled palmitic acid was reduced back to 1-hexadecanol. The authors state that their results are consistent with the presence of a "fatty alcohol cycle" in which hexadecanol is synthesized from palmitate via acyl-CoA and simultaneously oxidized back to free fatty acid. There is similar evidence that even longer chain fatty alcohols (C24–C34), such as the constituents of TOS, are metabolised by a similar fatty alcohol cycle in humans (Hargrove et al. 2004).

Block 11: Sterols and Steroids

The substances in this block, β-sitosterol, β-sitostanol, campesterol, lupeol, cycloartenol, citrostanienol, (α-sitosterol), methyl betulinate and betulin and are loosely referred to as phytosterols or phytosteroids.  Several phytosterols, which are ingested in significant amounts from the diet, have been shown to inhibit cholesterol uptake, and to some degree cholesterol biosynthesis, especially β-sitosterol, β-sitostanol and campesterol from the above list.  Consequently, these have been the subject of much research on their toxicokinetics, safety and biochemistry.  Where they have been tested, β-sitosterol, β-sitostanol, campesterol have poor bioavailability (poorer than the structurally similar cholesterol) in mammals, including humans.  They have a rapid initial rate of excretion but a much slower final rate.  Metabolism is slow and there is no evidence that phase II conjugates such as glucuronides are formed.  Little data could be found on the metabolism of lupeol, cycloartenol, citrostanienol (α-sitosterol) but betulin, and methyl betulinate/betulinic acid, which have been shown to have useful anti-cancer and anti-HIV activities were shown to be subject to extensive metabolism and phase II conjugation in rats. They occur primarily in Betula (birch) bark and wood, so their concentrations in TOS are likely to vary with the amount of birch wood used in pulping.

Several of the phytosterols (plant sterols) contained in TOS have been the subject of intense research scrutiny due to their effect on lowering the uptake of dietary cholesterol in humans.  The compounds that have been most studied from the group above are β-sitosterol, β-sitostanol and campesterol. They all have the same ABCD ring structure as cholesterol (although β-sitostanol is saturated at C4-C5) but the main differences from cholesterol is in the nature of the 17-substituent, which is branched at C25 with a methyl (campesterol) or ethyl (β-sitosterol, β-sitostanol. (For numbering of the sterol ring see Fig. 5.1.15). They all appear to work by displacing cholesterol from intestinal micelles and reduce the pool of absorbable cholesterol, but they are also rapidly taken up by enterocytes and increase expression of the ATP-binding cassette A1 sterol transporter. However, in comparison with cholesterol they are poorly absorbed, and an appreciable proportion of the material ingested is voided directly in the faeces (Ostlund, 2004).  Different phytosterols are absorbed to a different amount, for example β-sitostanol is adsorbed by the small intestine to a considerably lesser degree than β-sitosterol and this appears to be correlated with the effectiveness in lowering blood plasma cholesterol concentrations. viz. β-sitostanol is more effective than β-sitosterol (Becker et al. 1993, see below). Comparison of the toxicokinetics of the individual sterol components in TOS are to a degree complicated by the fact that many research papers have used mixtures of sterols (natural or artificial) in measuring the uptake and biological effect, so where possible work that has used single components has been used in this profile.

β-Sitosterol

β-sitosterol is an abundant phytosterol in nature, and is the sterol that is present in the highest concentration in TOS. The uptake of β-sitosterol (and β-sitostanol) were evaluated by Becker et al. (1993), who measured the absorption of the 14C labelled sterols in rats. The fates of [4-14C] β-sitosterol) and [4-14C] β-sitostanol were compared after oral or intravenous administration to rats. Excretion into the faeces of orally administered beta-sitostanol was significantly higher than that of β-sitosterol. More than 97% of β-sitostanol and 88% of β-sitosterol were recovered in the faeces within 7 days showing the relatively poor uptake of both compounds, however, the actual nature of the faecal metabolites was not established.  

The metabolic turnover, absolute oral bioavailability, clearance, and distribution volume of β-sitosterol were measured in humans by Duchateau et al. (2012).  [14C] β-Sitosterol was used as an isotopic tracer to distinguish pulse doses from dietary sources and was administered by both oral and intravenous routes.  The oral bioavailability was low (0.41%) and the turnover was 5.8 mg/day. Consequently, β-sitosterol was assessed as having a poor bioavailability in humans.

The bioavailability of β-sitosterol in beagle dogs was also assessed as being low.  When [3H]-labelled β-sitosterol (10 mg) was administered orally to beagle doges the absolute bioavailability upon was 9% with a mean residence time of about 185 h (Ritschel et al, 1990). The fates of [4-14C]-β-sitosterol and [4-14C]-β-sitostanol were compared after oral or intravenous administration to rats. Excretion into faeces of oral β-sitosterol was significantly higher than that of β-sitostanol. More than 97% of β-sitostanol and 88% of β-sitosterol were recovered in the faeces within 7 days. Thus, deposition of β-sitostanol was negligible in the tissues that were examined. Turnover in serum of sitostanol which was injected intravenously appeared to be more rapid than that of β-sitosterol; β-sitostanol was excreted as neutral steroids at a rate more than twice that of β-sitosterol. The rate of excretion of [3H]-cholesterol was slightly greater when β-sitostanol was administered simultaneously. The liver contained significantly less radioactivity after β-sitostanol than after β-sitosterol administration. More β-sitostanol than β-sitosterol was present in esterified form in serum and liver. The ratio of sterol in very low-density lipoprotein to that in high density lipoprotein was less for β-sitostanol or β-sitosterol than for endogenous cholesterol; this was particularly marked with β-sitostanol. These results suggest that β-sitostanol is a more effective hypocholesterolemic agent than β-sitosterol (Ikeda et al, 1978).

The fate of phytosterols in the rat was assessed by Sanders et al, (2000) as part of safety evaluation programme of phytosterols. Rats were dosed by oral gavage with 14C-labelled samples of cholesterol, β-sitosterol or β-sitostanol or 3H-labelled samples of β-sitostanol, campesterol, campestanol or stigmasterol dissolved in sunflower seed oil. Urine and faeces were collected for up to 96 hours after dosing. The amount of radiolabel remaining in the carcass was also measured.  The overall absorption of phytosterols was low as measured by tissue and carcass levels of radioactivity. Elimination from the body was mainly in the faeces and was initially very rapid, but traces of material were still being excreted at 4 days after dosing. Cholesterol was absorbed to the greatest extent (27% of the dose in females at 24 hours). Campesterol (13%) was absorbed more than β-sitosterol and stigmasterol (both 4%) which were absorbed more than β-sitostanol and campestanol (1-2%).

Phytosterols are susceptible to air-oxidation in the environment, for example, the analysis of baby food containing β-sitosterol showed the presence of 7-ketositosterol (García-Llatas et al, 2008). The extent of stigmasterol oxidation (2.9%) was even higher compared with β-sitosterol (1.4%). Consequently, the discovery of oxidised forms of phytosterols in blood plasma could be due either to absorption of oxidised metabolites or cytochrome-P450 oxidation in vivo. For example, the oxidised derivatives of phytosterols (oxyphytosterols) were identified in plasma samples from thirteen healthy human volunteers, using mass spectrometry (Grandgirard et al, 2004a). All the samples contained noticeable quantities of (24R)-5,beta,6,beta-epoxy-24-ethylcholestan-3beta-ol (β-epoxysitostanol) and (24R)-ethylcholestan-3,beta,5,alpha,6,beta-triol (sitostanetriol) and also trace levels of (24R)-5,alpha,6,alpha-epoxy-24-ethylcholestan-3beta-ol (alpha-epoxysitostanol), (24R)-methylcholestan-3beta,5alpha,6beta-triol (campestanetriol) and (24R)-ethylcholest-5-en-3beta-ol-7-one (7-ketositosterol). However, in a later paper (Grandgirard et al, 2004b) the authors showed that sitostanetriol was not formed in vivo from β-sitosterol in the rat and concluded that the identification of oxidised phytosterols in human subjects was to a degree artefactual and was due to the uptake of oxidised phytosterols in the diet.

The metabolic transformation of β-sitosterol in vitro to 26-hydroxy-β-sitosterol (Fig. 5.1.15, metabolite 1) and 29-hydroxy-β-sitosterol (Fig. 5.1.15, metabolite 2) using a recombinant form of the human cytochrome P450 CYP27A1 was shown by Ehrhardt et al. (2016).

The metabolism of phytosterols in rat faeces and liver microsomes was investigated. Faeces were collected after phytosterols (a well characterized mixture of beta-sitosterol 40%, campesterol 30% and dihydrobrassicasterol) were administered orally (0.5 g/kg) to rats. Metabolites of phytosterols were identified using GC/MS. Three peaks were eluted at 12.47, 12.65, 12.87 min and had characteristic molecular ions m/z 428, 430, 432, respectively. Three faecal metabolites were identified as 1,4-androstadiene-3,17-dione (Fig. 5.1.15, metabolite 5), 4-androstene-3,17-dione (Fig. 5.1.15 metabolite 4) and androstane-3,17-dione ((Fig. 5.1.15 metabolite 3). No metabolites could be detected in the rat liver microsomal reaction mixture. The results suggest that the metabolites of phytosterols in rat faeces are formed by oxidation at the 3-position, saturation at the 5- and 6- positions, and 17- side chain cleavage in the rat large intestine.  Thus, it is likely that the metabolites found in faeces are due to bacterial rather than mammalian metabolic transformations (see Sripalakit et al, 2006).

No evidence could be found for sitosterol (or any other phytosterol) phase II conjugate formation such as 3-glucuronides so this, together with the other data cited above, suggest they are rather refractory to metabolic transformation (catabolism) in mammals.

The route of degradative metabolism of β-sitosterol in rats (faecal/microbial) and humans (in vitro) is shown in Figure 5.1.15

 

Campesterol

Campesterol is a close structural analogue of β-sitosterol (25-methyl versus 25-ethyl analogue), thus its uptake and toxicokinetics closely follow its analogue. There is some evidence that intestinal absorption is greater than β-sitosterol. (see entry on β-sitosterol above and Sanders et al, 2000).  No separate information could be found on its catabolism but the structural similarity to β-sitosterol is so close, it would be expected to be similar.

β-Sitostanol

β-Sitostanol is also a close structural analogue of β-sitosterol with the 5C=C6 double bond in the B ring reduced to 5CC6 (hence “stanol” rather than “sterol”).  Thus, its uptake and toxicokinetics closely follow its analogue, although its intestinal absorption is considerably less than β-sitosterol and deposition of β-sitostanol was negligible in the rat tissues that were examined by Ikeda et al. (1978). (see entry on β-sitosterol above and Sanders et al, 2000 and particularly Ikeda et al, 1978).  No separate information could be found on its catabolism but the structural similarity to β-sitosterol is so close, it would be expected to be similar.

Lupeol

Lupeol is a natural triterpenoid found in many plant species such as mango and it is the principal active component of many traditional herbal medicines. No information on the mammalian metabolism of lupeol could be found in the literature; however, in a paper to assess the inhibition and induction potential of lupeol and betulin on cytochromes P450 (CYP)1A2, CYP2C11, CYP2D6 and CYP3A2 activities in rat liver microsomes both compounds were inactive (Seervi et al, 2016).

Cycloartenol

Cycloartenol is an alternative biosynthetic intermediate to lanosterol in the plant biosynthesis pathway to phytosterols (Ohyama et al, 2011).  No information on the mammalian degradative metabolism of cycloartenol could be found in the literature.

Citrostanienol (α-Sitosterol)

No information on the metabolism of citrostanienol/α-sitosterol could be found in the literature.

Betulin

Betulin is an active natural pentacyclic triterpene compound with anti-cancer and anti-HIV activities. It is found in birch bark and wood and related trees such as Alnus (alder).  No information on the mammalian uptake of betulin could be found in the literature; however, two papers have assessed its mammalian metabolic transformation.  In a recent paper, Zhang et al. (2018) used a highly efficient method to screen and identify the metabolites and to assess the metabolic profiles of betulin in vivo using UHPLC-Q-TOF-MS/MS system based on multiple mass defect filter data acquisition (MMDF) and multiple data processing techniques. 56 phase I and 6 phase II metabolites were detected in rats after oral administration of betulin. The main biotransformation routes of betulin were identified as demethylation, dehydroxylation, deoxidization, dehydration. Conjugation with sulfate, taurine, cysteine and N-acetylcysteine groups produced 6 phase II metabolites.  Since betulin is found at levels only just above the 0.1% cut-off for this profile, a detailed metabolic pathway for betulin is not presented in this profile.

In an even more recent in vitro study, the glucuronidation and sulfation of betulin in human and rat liver microsomes and cytosol were tested (Hu et al. 2019). One betulin glucuronide metabolite was observed after incubation with human and rat liver microsomes and two betulin sulfate metabolites were found in human and rat liver cytosols although the chemical structure of these two conjugates were not determined. These results suggest that glucuronidation and sulfation are important phase II catabolic pathways for betulin. The structure of betulin is shown in Fig. 5.1.16, together with the two hydroxyl groups (at C3 and C28) that are available for conjugation with glucuronic acid or sulfate (red).

Methylbetulinate and Betulinic acid

These are also found in high concentrations in birch wood and are under study for similar pharmaceutical applications as betulin.  Zhang et al. (2019) used the same methodology to study the degradation of betulinic acid as the same group had used with betulin and detected 46 metabolites that were structurally characterised and indicated a similar catabolic pathway to betulin.

Block 12: Lignin, cellulose and fibre

Not assessed due to difficulty in measuring the metabolism of polymers in vivo.

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