Docosahexaenoic acid is both a product of and a precursor to tetracosahexaenoic acid in the rat

Abstract

Tetracosahexaeoic acid (THA; 24:6n-3) is thought to be the immediate precursor of DHA in rodents; however, the relationship between THA and DHA metabolism has not been assessed in vivo. Here, we infused unesterified ²H₅-THA and ¹³C₂₂-DHA, at a steady state, into two groups of male Long-Evans rats and determined the synthesis-secretion kinetics, including daily synthesis-secretion rates of all 20-24 carbon n-3 PUFAs. We determined that the synthesis-secretion coefficient (a measure of the capacity to synthesize a given fatty acid) for the synthesis of DHA from plasma unesterified THA to be 134-fold higher than for THA from DHA. However, when considering the significantly higher endogenous plasma unesterified DHA pool, the daily synthesis-secretion rates were only 7-fold higher for DHA synthesis from THA (96.3 ± 31.3 nmol/d) compared with that for THA synthesis from DHA (11.4 ± 4.1 nmol/d). Furthermore, plasma unesterified THA was converted to DHA and secreted into the plasma at a 2.5-fold faster rate than remaining as THA itself (26.2 ± 6.3 nmol/d), supporting THA’s primary role as a precursor to DHA. In conclusion, using a 3 h infusion model in rats, we demonstrate for the first time in vivo that DHA is both a product and a precursor to THA.

Tetracosahexaenoic acid (THA; 24:6n-3) was first reported to be the immediate precursor to DHA in 1991 by Voss et al. (1), where n-3 docosapentaenoic acid (DPAn-3; 22:5n-3) had previously been thought to be directly converted to DHA via a Δ4-desaturation reaction. Later, a lack of labeled DHA accumulation in Zellweger’s patients (2, 3) signaled a role for the peroxisomes in the β-oxidation of THA to DHA, following the elongation and subsequent Δ6-desaturation of DPAn-3 to n-3 tetracosapentaenoic acid (TPAn-3; 22:5n-3) and THA, respectively. Termed the “Sprecher pathway” (DPAn-3 → TPAn-3 → THA → DHA), this series of reactions have since been accepted as vital steps in the synthesis of DHA; however, recent evidence from human cells is contradictory and suggests that THA is a product, not a precursor, of DHA (4, 5). Currently, it appears possible that rats and humans synthesize DHA through different pathways; however, often-overlooked data from 1991 show elongation of DHA to THA in addition to the Δ6-desaturation TPAn-3 and β-oxidation of THA to DHA (1). Furthermore, the human cell study demonstrated that in the absence of functional Δ6-desaturase activity, labeled DPAn-3 could be converted directly to DHA (5); however, it is possible that this Δ4-desaturase activity is upregulated in response to the absence of a Δ6-desaturase enzyme.

Recently, our laboratory demonstrated the sensitivity by GC/MS to determine daily synthesis-secretion rates for all n-6 and n-3 PUFAs following 3 h steady-state infusions of linoleic acid (18:2n-6) and α-linolenic acid (ALA; 18:3n-3), respectively, in Long-Evans rats (6, 7). From these studies, we determined that the synthesis-secretion rates of THA and DHA from plasma unesterified ALA (7) and from plasma unesterified ALA and EPA in response to DHA feeding (6) are remarkably similar, despite much higher plasma levels of DHA. In addition, animals fed a high-ALA diet resulted in higher plasma THA levels compared with a low-ALA diet, but animals fed a high- versus low-DHA diet showed no change in plasma THA, suggesting that THA is indeed a precursor to DHA (8). These studies suggest some potentially unique metabolic differences between DHA and THA, despite their close relationship in the biosynthetic pathway. The ability to determine synthesis-secretion rates of the least abundant n-3 PUFA in living animals lends to the possibility of performing 3 h infusions with these low-abundance molecules to determine actual rates of synthesis between THA and DHA.

In the present study, we perform 3 h steady-state infusions of ²H₅-THA and ¹³C₂₂-DHA into Long-Evans rats by the method developed by Rapoport et al. (9) and modified by our laboratory (10). This is the first study to infuse a THA or DHA tracer into a living animal to determine synthesis rates of DHA and THA, respectively. Briefly, we determined that plasma unesterified THA is converted to DHA, but also demonstrated that plasma unesterified DHA is converted to THA with the predominant rate of the reaction being for DHA synthesis from THA. In addition, we showed that THA is preferentially converted to DHA compared with acylation into complex lipids, a novel finding compared with other n-3 and n-6 PUFA assessed via the same animal models.

MATERIALS AND METHODS

Animals

All experimental procedures were performed in agreement with the policies set out by the Canadian Council on Animal Care and were approved by the Animal Ethics Committee at the University of Toronto. Eight 18-day-old male nonlittermate Long-Evans pups with dam were ordered from Charles River Laboratories (St. Constant, QC, Canada). Following arrival at the University of Toronto, the dam and pups were acclimated for 3 days and then weaned at 21 days old. The dams were placed on a standard chow diet (Teklad 2918, Harlan, Madison, WI) immediately upon arriving at the University of Toronto, and the pups were placed on the same diet for 11–12 weeks following weaning until 14–15 weeks of age. During this time and prior to cannulation, rats were handled frequently and housed in pairs (7).

Diets

The diet contained 6.2% fat, 18.6% protein, and 44.2% carbohydrates by weight. The fatty acid composition (% weight in total fatty acids) was determined by gas chromatography-flame ionization detection (GC/FID) to be 6.17 ± 0.01% (n = 3, means ± SEM) ALA, 11.83 ± 0.04% 16:0, 3.18 ± 0.001% 18:0, 1.09 ± 0.001% 18:1n-7, 19.97 ± 0.02% 18:1n-9, and 53.53 ± 0.08% 18:2n-6, equaling more than 95% of all fatty acids in the diet.

Surgery and ²H₅-THA, ¹³C₂₂-DHA, and ²H₅-ALA infusion

At 11–12 weeks postweaning, 14–15-week-old rats were subjected to surgery to implant a catheter into the jugular vein and the carotid artery, as previously described in detail (10). Baseline plasma unesterified and total fatty acid concentrations were measured in plasma and drawn from the carotid artery 1 day after surgery and 2 days prior to stable isotope infusion. Modified from the method of Rapoport et al. (9), the first group of animals (n = 4) was infused into the jugular vein with 0.070 μmol/100 g body weight of ²H₅-THA (purity >95% confirmed by GC-FID and GC/MS; n = 3; Cayman Chemical Co., Ann Arbor, MI). Due to the higher endogenous unesterified plasma ALA and DHA pools, significantly more tracer was used to try to better match these endogenous levels. The second group of animals (n = 4) was infused with 0.843 μmol/100 g body weight of ¹³C₂₂-DHA (Cambridge Isotope Laboratories, Inc., Tewksbury, MA). A single rodent from the second group of animals was coinfused with 0.843 μmol/100 g body weight of ²H₅-ALA (Cambridge Isotope Laboratories) to compare with previous reports as a positive control to confirm that the infusions were being administered properly. Infusate preparation, 3 h steady-state infusions, and blood collections from the carotid artery (0, 30, 60, 90, 120, 150, and 180 min) were performed as previously described in detail (10). All blood samples were centrifuged for 15 min at 500 g (PC-100 microcentrifuge; Diamed, ON, Canada) under refrigeration, and the serum was collected and stored at −80°C.

Tracer purity determination

At the end of a 3 h infusion, 10 μl aliquots of the ²H₅-THA infusate and 125 μl aliquots of the ²H₅-ALA + ¹³C₂₂-DHA (n = 3) infusate were collected and extracted by a method modified from Folch, Lees, and Sloane Stanley (11). Briefly, lipids were extracted with 2:1:0.75 chloroform:methanol:0.88% potassium chloride (vol:vol:vol) containing 5 μg of heptadecanoic acid (17:0; NuChek Prep Inc., Elysian, MN) as internal standard. The mixtures were vortexed and centrifuged at 500 g for 10 min, and the lower, chloroform lipid-containing layer was pipetted into a new test tube. The total lipid extract (TLE) was then dried down under a stream of nitrogen, hydrolyzed, derivatized to pentafluorylbenzyl (PFB) esters, and run on the GC/MS for identification of any contaminant tracers that would yield false positives for metabolism of infused tracers. For example, in the ²H₅-ALA + ¹³C₂₂-DHA infusate, it was important to confirm the absence of other ¹³C₂₂-n-3 PUFAs (i.e., ¹³C₂₂-DPAn-3, ¹³C₂₂-TPAn-3, and ¹³C₂₂-THA) to confirm that all ¹³C₂₂-n-3 PUFAs were derived from the metabolism of ¹³C₂₂-DHA.

Determination of plasma volume

Plasma volume was determined using the method of Schreihofer, Hair and Stepp and modified by our lab (12, 13). Briefly, a known amount of Evans Blue dye was injected into the jugular vein of a separate group of rats (n = 3). Fifteen minutes following injection, 1 ml blood was drawn from the carotid artery, twice. The plasma was collected as described above, and 100 μl of plasma was diluted into 1 ml of saline. Absorbance of plasma in saline was determined at 604 nm with a Nanodrop 1000 and compared with a standard curve, and the concentration of the Evans Blue dye was determined. Concentration of the dye was then used to determine plasma volume.

Lipid extractions from baseline serum

TLEs were obtained from 50 μl of serum by the extraction method described earlier. Aliquots of one-fifth of the TLE for total fatty acids and four-fifths for unesterified fatty acid determinations were taken. TLEs for total fatty acid determinations were stored for hydrolysis as described later. TLEs for unesterified fatty acid determinations were evaporated under N₂ gas and reconstituted in 50 μl chloroform, plated on a silica 60 G plate (Merck Millipore, Billerica, MA), and developed using TLC with 60:40:2 heptane:diethyl ether:glacial acetic acid for separation of unesterified fatty acids, as previously described (14). Unesterified fatty acidsFAs were scraped from TLC plates and extracted from the silica by the method described earlier. Isolated unesterified fatty acids were stored until derivatization to PFB esters.

Lipid extractions from stable isotope-infused plasma

TLE were obtained from 80 μl of stable isotope-infused plasma by the Folch method as described earlier containing 5 μg of heptadecanoic acid (NuChek Prep Inc.) as internal standard. Three-quarters of the TLE was stored for hydrolysis and stable isotope enrichment of total fatty acids, and the remaining one-quarter of the TLE was used for isolation of isotopically labeled unesterified fatty acid by the TLC method described earlier. The isolated labeled unesterified fatty acids were stored for later derivatization to PFB esters.

Hydrolysis of total endogenous and isotopically labeled total fatty acid pools

The TLE from baseline serum and infused plasma used to determine endogenous and labeled fatty acid concentrations, respectively, was evaporated under nitrogen, and the lipids were hydrolyzed in 2 ml of 10% potassium hydroxide in methanol (wt:vol), as previously described (8, 14, 15). Hydrolyzed fatty acids from the total lipid pool were collected and stored at −80°C for later derivatization to PFB esters.

PFB esterification

The plasma unesterified fatty acids and hydrolyzed fatty acids from total lipid pools were dried under nitrogen and derivatized to PFB esters, as previously described and modified by our lab (7, 15). Briefly, 100 μl of acetonitrile:diisoproplyamine:PFB bromide (1,000:10:1, vol:vol:vol) was added and heated at 60°C for 15 min. The reagents were then evaporated under nitrogen, realiquoted in 40 μl of hexane, and analyzed for endogenous and labeled fatty acids by GC/MS.

Endogenous and labeled fatty acid determination by GC/MS

Fatty acid PFB esters were analyzed on an Agilent 5977A quadrupole mass spectrometer coupled to an Agilent 7890B gas chromatograph (Agilent Technologies, Mississauga, ON, Canada) in negative chemical ionization mode, as described previously (15) and recently modified by our lab (8). The fatty acid PFB esters were injected via an Agilent 7693 autosampler into a DB-FFAP 30 m × 0.25 mm inner diameter × 25 μm film thickness capillary column (J&W Scientific from Agilent Technologies) interfaced directly into the ion source with helium as the carrier gas and 5.0 methane (Praxair Canada Inc., Toronto, ON, Canada) as the ionization gas. Fatty acids were analyzed in selected ion-monitoring mode using [M-H] for parent ion identification with ion dwell times of 500 ms. The [M-H] parent ions (m/z) for endogenous and labeled fatty acids are as follows for unlabeled PUFAs, ²H₅-PUFA and ¹³C_20-22, respectively: 277.3 and 282.3 for ALA; 301.3, 306.3, and 321.3 for EPA; 329.3, 334.3, 349.3, and 351.3 for DPAn-3; 327.3, 332.3, and 349.3 for DHA; 357.3, 362.3, and 379.3 for TPAn-3; and 355.3, 360.3, and 377.3 for THA.

Equations

To determine the synthesis rates of n-3 PUFAs from THA and DHA, the appearance of ²H₅-PUFA and ¹³C₂₂-PUFA, respectively, in the plasma-esterified pool were measured and fit to a Boltzmann sigmoidal curve ([²H₅/¹³C₂₂-PUFA] × plasma volume vs. time) using nonlinear regression (12) (GraphPad Prism Version 4.0, La Jolla, CA). In addition, the retroconversion of ¹³C₂₂-DHA to ¹³C₂₀-EPA and ¹³C₂₀-DPAn-3 was included in our calculations. At any point on the curves, the slope (S) is determined by the ability of the body to synthesize ²H₅-PUFA from ²H₅-THA or ¹³C_20-22-PUFA from ¹³C₂₂-DHA and the ability of the periphery to uptake ²H₅-PUFA (equation 1) or ¹³C_20-22-PUFA (equation 2), respectively.

$$S=k_{1,\mathrm{PUFA}}{\left[{}^2\mathrm{H}_5-\mathrm{THA}\right]}_{\mathrm{unest}}-k_{-1,\mathrm{PUFA}}{\left[{}^2\mathrm{H}_5-\mathrm{PUFA}\right]}_{\mathrm{est}}$$ (Eq. 1)

$$S=k_{1,\mathrm{PUFA}}{[{}^{13}C_{22}-\mathrm{DHA}]}_{\mathrm{unest}}-k_{-1,\mathrm{PUFA}}{[{}^{13}C_{20-22}-\mathrm{PUFA}]}_{\mathrm{est}}$$ (Eq. 2)

where k_1,PUFA is the steady-state synthesis-secretion coefficient for any PUFA, representing a measure for the amount of infused label that is converted to downstream products; [²H₅-THA]_unesterified, [¹³C₂₂-DHA]_unesterified are the plasma concentrations of the infusate; k_-1,PUFA is the disappearance coefficient for any individual PUFA; and [²H₅/¹³C_20-22-PUFA]_esterified is the concentration of an individual PUFA in the plasma that has been synthesized from the respective infusate, packaged into lipoprotein, and secreted into the plasma.

The maximum first derivative (S_max) of the curves is assumed to be the time point when the uptake of esterified labeled PUFA from the periphery is negligible, i.e., 0 (equations 3 and 4).

$$S_{\max }=k_{1,\text{PUFA}}{\left[{}^2\mathrm{H}_5-\text{ALA}\right]}_{\text{unest}}-0$$ (Eq. 3)

$$S_{\max }=k_{1,\text{PUFA}}{\left[{}^{13}\mathrm{C}_{22}-\text{DHA}\right]}_{\text{unest}}-0$$ (Eq. 4)

Therefore, the derivative at this point is equal to the rate of ²H₅-PUFA or ¹³C_20-22-PUFA synthesis secretion. By correcting the S_max by the substrate tracee:tracer ratio, the rates of actual PUFA synthesis, J_syn,PUFA (nmol/min) (12), are determined (equations 5 and 6).

$$J_{syn,\text{PUFA}}=S_{\max }\left({{\left[\text{ALA}\right]}_{\text{unest}}/\left[{}^2\mathrm{H}_5-\text{ALA}\right]}_{\text{unest}}\right)=k_{1,\text{DHA}}{\left[\text{ALA}\right]}_{\mathrm{unest}}$$ (Eq. 5)

$$J_{syn,\text{PUFA}}=S_{\max }\left({{\left[\text{DHA}\right]}_{\text{unest}}/\left[{}^{13}\mathrm{C}_{22}-\text{DHA}\right]}_{\text{unest}}\right)=k_{1,\text{DHA}}{\left[\text{DHA}\right]}_{\text{unest}}$$ (Eq. 6)

The diet consumed for this study contained no n-3 PUFA other than ALA; therefore it can be assumed that the plasma total PUFAs were constant during the infusion period. Therefore, the turnover rate (F_PUFA) and half-life (t_1/2,PUFA) of total PUFA in the plasma can be determined by equations 7 and 8, respectively.

$${\mathrm{F}}_{\text{PUFA}}={\mathrm{J}}_{\text{PUFA}}/\left({\mathrm{V}}_{\text{plasma}}*{\left[\text{PUFA}\right]}_{\text{esterified}}\right)$$ (Eq. 7)

$${\mathrm{t}}_{\text{1}/2,\text{PUFA}}=0.693/{\text{F}}_{\mathrm{PUFA}}$$ (Eq. 8)

Statistics

Differences in n-3 PUFA synthesis rates of downstream products from an individual tracer (²H₅-THA or ¹³C₂₂-DHA) were assessed by one-way ANOVA followed by Tukey’s honest significant difference posthoc test. Student’s t-test was used to compare kinetic parameters of an individual n-3 PUFA product between the two tracers (i.e., ²H₅-THA vs. ¹³C₂₂-THA). Statistical significance was set at P < 0.05 for all tests. Normality was assessed by the Shapiro-Wilk test for normality, and in the case of nonnormally distributed groups, the data were log-transformed prior to statistical analysis. All data are presented as mean ± SEM.

RESULTS

Tracer purity

The ²H₅-THA infusate and the ²H₅-ALA + ¹³C₂₂-DHA infusate were assessed by GC/MS for contaminant isotopes that would result in false-positive determinations of the origin of n-3 PUFA metabolites, with representative chromatograms presented (Fig. 1). An external reference standard chromatogram is also presented (Fig. 1A). In the ²H₅-ALA + ¹³C₂₂-DHA infusate (Fig. 1B), no ¹³C_20-22-n-3 PUFA contaminants were detected for EPA, DPAn-3, TPAn-3, or THA, with ¹³C₂₂-DHA being the only ¹³C-n-3 PUFA identified. In the ²H₅-THA infusate (Fig. 1C), no ²H₅-n-3 PUFA contaminants were detected for ALA, EPA, DPAn-3, DHA, or TPAn-3. The only n-3-PUFA isotope detected was ²H₅-THA.

Fig. 1.

Representative GC/MS chromatograms for external standard (A), ²H₅-ALA + ¹³C₂₂-DHA infusate (B), and ²H₅-THA infusate (C). n.d., not detected.

Plasma-labeled and unlabeled n-3 PUFA concentrations

Plasma unesterified ALA, DHA, and THA (Fig. 2A), unesterified ²H₅-ALA, ¹³C₂₂-DHA, and ²H₅-THA (Fig. 2B), and tracer/tracee ratios (²H₅-ALA/ALA, ¹³C₂₂-DHA/DHA, and ²H₅-THA/THA) (Fig. 2C) were determined and are presented in Fig. 2. For ALA, DHA, and THA, respectively, endogenous unesterified fatty acid concentrations (nmol/ml ± SEM) were 4.4 ± 0.7, 2.8 ± 1.0, and 0.12 ± 0.03; labeled unesterified fatty acid infusates were 0.30 ± 0.06, 0.41 ± 0.12, and 0.052 ± 0.004; and tracer/tracee ratios were 0.08 ± 0.02, 0.17 ± 0.04, and 0.52 ± 0.12, respectively.

Fig. 2.

Baseline plasma unesterified fatty acid concentrations (A), unesterified labeled fatty acid infusate concentrations (B), and tracer/tracee ratios for infusate of n-3 PUFA (C) in male Long-Evans rats. Values are expressed as mean ± SEM, n = 4.

Total plasma n-3 PUFA concentrations (nmol/ml ± SEM; data not shown) were also determined for turnover and half-life calculations in the ²H₅-THA and the ²H₅-ALA + ¹³C₂₂-DHA infused animals (data not shown). In the ²H₅-THA infused animals, total EPA, DPAn-3, DHA, and THA were 17.8 ± 3.5, 20.9 ± 4.1, 126 ± 31, and 1.88 ± 0.49, respectively, and in the ¹³C₂₂-DHA, total concentrations were 9.1 ± 0.8, 11.0 ± 1.7, 68.5 ± 14.4, and 0.84 ± 0.13, respectively. No significant differences (P > 0.05) were determined between the two infusion groups for any n-3 PUFA.

n-3 PUFA synthesis-secretion coefficients, rates, turnovers, and half-lives

Representative infusion curves, S_max, and representative GC/MS chromatograms for the synthesis secretion of THA and DHA from plasma unesterified DHA (Fig. 3A) and THA (Fig. 3B) are shown. Furthermore, a statistical comparison for the synthesis-secretion kinetic parameters of DHA from plasma unesterified THA to the synthesis secretion of THA from plasma unesterified DHA is presented (Table 1). Most strikingly, the synthesis-secretion coefficient for DHA from THA (0.539 ± 0.098, ml/min ± SEM) is 134-fold higher (P < 0.01) than for THA from DHA (0.004 ± 0.002); however, when determining daily synthesis-secretion rates (96 ± 31 vs. 11.4 ± 4.1, nmol/day ± SEM), this difference (P = 0.036) is reduced to only 7-fold. Similarly, the turnover and half-life of newly synthesized plasma DHA from unesterified THA is 18-fold lower (P < 0.001) and 15-fold higher (P < 0.0001), respectively, compared with newly synthesized THA from unesterified DHA.

Fig. 3.

Representative infusion curves, maximum first derivatives, and representative GC/MS chromatograms for DHA and THA products of ¹³C₂₂-DHA (A) and ²H₅-THA (B).

TABLE 1.

Kinetic parameters for the whole-body synthesis secretion of n-3 PUFAs following infusion of ¹³C₂₂-DHA or ²H₅-THA

Source	N-3 PUFA	Maximum First Derivative (nmol/min)	Synthesis-Secretion Coefficient (ml/min)	Daily Synthesis-Secretion Rate (nmol/day)	Turnover (turnover/day)	Half-life (days)
DHA	EPA	0.016 ± 0.003a	0.058 ± 0.022a	179 ± 61a	2.40 ± 0.66a	0.390 ± 0.136a
	DPAn-3	0.006 ± 0.002b	0.020 ± 0.010a	50.9 ± 10.7a	0.652 ± 0.165b	1.28 ± 0.29b
	TPAn-3	n.d.	n.d.	n.d.	n.d.	n.d.
	THA	0.001 ± 0.0002c	0.004 ± 0.002a	11.4 ± 4.1a	1.74 ± 0.48ab	0.468 ± 0.085a
	DHA	0.397 ± 0.076d	1.32 ± 0.45b	4434 ± 1575b	8.17 ± 2.28c	0.102 ± 0.022a
THA	EPA	0.001 ± 0.0002*,a	0.016 ± 0.003a	2.88 ± 0.89*,a	0.019 ± 0.004*,a	42.4 ± 10.9*,a
	DPAn-3	0.001 ± 0.0003*,a	0.016 ± 0.004a	2.75 ± 0.78*,a	0.016 ± 0.003*,a	48.8 ± 9.3*,a
	TPAn-3	n.d.	n.d.	n.d.	n.d.	n.d.
	THA	0.008 ± 0.002*,b	0.157 ± 0.024*,a	26.2 ± 6.3a	1.80 ± 0.16b	0.394 ± 0.037b
	DHA	0.027 ± 0.005*,#,c	0.539 ± 0.098#,b	96.3 ± 31.3*,#,b	0.093 ± 0.007*,#,a	7.55 ± 0.61*,#,b

Different superscript letters represent statistically different kinetic values between fatty acids and within a kinetic measure following a significant one-way ANOVA and Tukey’s posthoc test. Values are expressed as means ± SEM, n = 4. n.d., not detected.

* P < 0.05 [significantly different kinetic values between sources (THA vs. DHA) for a specific n-3 PUFA as determined by Student’s t-test].

^#P < 0.05 (significantly different kinetic values for THA synthesis from DHA vs. DHA synthesis from THA as determined by Student’s t-test).

A comprehensive assessment of synthesis-secretion kinetic parameters for all plasma n-3 PUFAs from plasma unesterified DHA and THA is presented (Table 1). In addition, synthesis-secretion kinetics from plasma unesterified ALA for two animals are presented as a positive control to demonstrate that the pathway is behaving within an acceptable range (supplemental Table S1). Most interestingly, the synthesis-secretion coefficient of total plasma DHA from plasma unesterified DHA compared with THA is not different (P > 0.05), despite a 45-fold difference (P < 0.001) in the daily synthesis-secretion rates. Furthermore, the synthesis-secretion coefficient and the daily synthesis-secretion rate for the synthesis of DHA from THA is 2- and 3-fold (P < 0.05) higher, respectively, compared with the synthesis of THA itself. In addition, the turnover of all measured n-3 PUFA was faster (P < 0.05) when from plasma unesterified DHA, except THA, which was not different (P > 0.05) between the two plasma sources. Finally, retroconversion of both DHA and THA to EPA and DPAn-3 were detected; however, these daily synthesis-secretion rates from DHA compared with THA were 60-fold and 18-fold higher (P < 0.05) for EPA and DPAn-3, respectively, with no differences (P > 0.05) in the synthesis-secretion coefficient. Importantly, labeled DPAn-3 was determined to be ¹³C₂₀-DPAn-3 most likely via elongation of newly formed EPA, and direct conversion of DHA to DPAn-3 via the appearance of ¹³C₂₂-DPAn-3 could not be detected (data not shown).

DISCUSSION

In the current study, we have for the first time using living animals through the infusion of ²H₅-THA over 3 h demonstrated that DHA is the primary n-3 PUFA product of THA. However, we have also shown for the first time in vivo the elongation of ¹³C₂₂-DHA back to THA in Long-Evans rats, and a modified n-3 PUFA biosynthesis pathway is presented in Fig. 4. The predominant rate of flow between these two n-3 PUFAs is from THA to DHA compared with DHA to THA. Importantly, the high purity of our tracers strengthens our conclusions with the knowledge that the only source of labeled DHA could be THA and vice versa. Surprising to many might be that the hallmark study by Voss et al. (1) in 1991, which was the first to demonstrate the need for a second Δ6-desaturation followed by peroxisomal β-oxidation to DHA, demonstrated that DHA can be elongated in isolated rat liver microsomes. They showed that a small proportion of 75 μM ¹⁴C-DHA is chain elongated to THA, and, when compared with acylation of DHA into phospholipids, DHA elongation was insignificant. Their conclusions were reasonable based on the labeled precursors and products in isolation; however, our 3 h infusion model (12) determines daily synthesis-secretion rates by correcting the labeled precursor-product ratio (synthesis-secretion coefficient) by the endogenous unesterified PUFA precursor pool. Significantly, this correction narrows the difference between the two rates of travel from 134-fold when considering the tracers only to merely 7-fold for the daily synthesis-secretion rate. Furthermore, because the liver DHA-CoA pool might dilute the labeled DHA more than the THA-CoA would dilute the labeled THA (14), the THA synthesis-secretion rates are expected to be closer to the DHA synthesis-secretion rates. The elongation of DHA to THA is likely mediated by the elongation of very-long-chain 2 (Elovl2) enzyme that has a high affinity for 22C-PUFA (16–18), particularly in comparison to Elovl5 that has very low affinity for 22C-PUFA and high affinity for 20C-PUFA (16).

Fig. 4.

n-3 PUFA biosynthesis pathway determined in 1991 compared with in 2018: Δ6-desaturation, elongation, and Δ5-desaturation converting ALA to EPA (1); elongation of EPA to DPAn-3 (2); elongation of DPAn-3 to TPAn-3 (3); Δ6-desaturation of TPAn-3 to THA (4); previously discovered peroxisomal β-oxidation of THA to DHA in 1991 (5); retroconversion of DHA to EPA (6); theoretical Δ4-desaturation of DPAn-3 to DHA reported in humans (7); and newly reported elongation of DHA to THA in the present 2018 study (8).

Our animals were fed a standard chow diet containing ALA as the only n-3 PUFA present. The inclusion of other n-3 PUFAs in the diet, particularly DHA, may result in even higher rates of THA synthesis secretion from plasma-unesterified DHA. Previously, we fed animals a DHA-free diet or a 2% DHA in total fat diet for 8 weeks and found that serum THA levels did not change, whereas serum DHA levels were significantly higher when fed the 2% DHA diet (6, 8). Furthermore, plasma unesterified DHA increased 12-fold and THA only 2-fold in response to a 2% DHA diet (6); however, these data were not reported in our previous study. This suggests that when animals are fed DHA, the higher plasma unesterified DHA would result in daily THA synthesis-secretion rates that are similar to the rates determined for DHA from plasma unesterified THA. However, because our study is the very first to infuse labeled THA and labeled DHA into free-living animals for the determination of daily synthesis-secretion rates to DHA and THA, respectively, the effect of DHA feeding on these metabolic questions has never been tested, and future studies need to be performed to address these unknowns.

What we cannot conclude based on our study design and results is the presence (or absence) of a Δ4-desaturase step that directly desaturates DPAn-3 to DHA while bypassing the Sprecher pathway. As a result, we also cannot clearly state what the predominant precursor to THA is, DHA or TPAn-3. It is possible that the n-3 PUFA biosynthetic pathway behaves very differently in isolated liver microsomes or hepatocytes compared with in living mammals. In such a scenario, DPAn-3 could be desaturated directly to DHA by a Δ4-desaturase as evidenced in human cells (4, 5), further elongated to THA, and then quickly β-oxidized back to DHA in the peroxisome. Interestingly, cells grown in the presence of labeled DHA demonstrated an up to 5-fold higher retroconversion of DHA to EPA compared with elongation to THA (4). We showed a similar, albeit larger (15-fold), rate of retroconversion over elongation that may continue to suggest similarities between rodent and human models of n-3 PUFA metabolism. Although the bulk of the evidence in rodents supports the Sprecher pathway, future studies could aim to clarify remaining questions relating to the final steps in DHA synthesis. For instance, 3 h infusions of DPAn-3, TPAn-3, THA, and/or DHA tracers in combination with in vivo inhibition of peroxisomal β-oxidation (19, 20), Δ6-desaturase inhibition (21, 22), or knockout (23) would provide a wealth of additional information and has the potential for a more complete understanding of these steps in the pathway. This could now be explored even further in humans, as a recent study has validated the infusion method in two individuals (24).

Interestingly, the rate of DHA synthesis from THA (96.3 ± 31.3 nmol/d) is very similar to our previously reported synthesis rates of DHA from ALA (47–157 nmol/d) (6, 7, 14, 25) and DHA from EPA (70–172 nmol/d) (6, 14) when feeding ALA as the only n-3 PUFA in the diet. In addition, these previously reported values are comparable, albeit somewhat lower, to the values presented in supplemental Table S1 and may be representative of the different diet used here. This further supports the recent conclusion from these studies that DHA synthesis rates are not as low as previously thought when considering ALA as the only substrate. DHA synthesis rates have frequently been reported to be very low and have been reviewed by our laboratory (26); however, if we consider for simplicity that DHA synthesis from ALA is 100 nmol/d, from EPA is 100 nmol/d, and from THA is 100 nmol/d, then DHA synthesis rates are at least three times higher than originally believed due to the inclusion of multiple DHA precursors. Furthermore, it is plausible that when we consider the four additional DHA precursors (18:4n-3, 20:4n-3, 24:5n-3, and, in particular, DPAn-3) that DHA whole-body synthesis rates will be even higher, and additional studies should be performed to test this.

Plasma unesterified THA in the rodent is converted to DHA at a faster rate than remaining as THA. This is shown very clearly, as the daily synthesis-secretion rate for DHA from THA is approximately two-and-a-half-fold higher than for THA remaining as itself and acylated into more complex lipids. This is a novel finding when compared with all previous 3 h infusion studies from our laboratory, as each of ALA, EPA, DHA, linoleic acid (18:2n-6), and DPAn-6 (22:5n-6) (6, 7, 10, 12, 14, 25) were more rapidly acylated into complex lipids versus converted to other PUFAs. This is likely related to the very low and relatively stable plasma levels of THA seen in the rodents (8), suggesting that THA must be rapidly metabolized in order to remain at these levels. This is also supportive of the original conclusion of Voss et al. (27) that the role of both THA and TPAn-3 are as “obligatory intermediates in 22:6(n-3) (DHA) biosynthesis” (1), and it has been shown in isolated peroxisomes + microsomes that β-oxidation of THA to DHA predominates over THA acylation into complex lipids. Unesterified DHA is also rapidly acylated into more complex lipids at a rate that is 390-fold higher than for elongation to THA, further supporting a molecular requirement for the presence of DHA in the body. This seemingly required conversion of THA into DHA is also highlighted by the lower relative rates of THA compared with DHA retroconversion to EPA and DPAn-3, indicating a likely requirement for the conversion of THA to DHA prior to retroconversion (28). Previous studies have also demonstrated DHA retroconversion to both EPA and DPAn-3 (29–31); however, they were unable to determine whether the DHA was converted directly to DPAn-3 with the removal of a single double bond or whether DHA was converted to EPA prior to elongation to DPAn-3. To the best of our knowledge, we show for the first time that retroconversion to EPA is likely to occur first and is supportive of the idea that the metabolic conversion of THA to DHA to EPA does not include a DPAn-3 intermediate (28). However, the low rates of DHA elongation to THA compared with retroconversion are not necessarily indicative of an unimportant role for THA, and the low conversion rates and plasma levels of THA could be further explained by rapid entry into other metabolic pathways. As such, future studies should be designed to investigate additional metabolic products of THA such as fatty acid ethanolamides or novel lipid mediators. Recently, a new class of neuroprotective lipid mediators, elovanoids, derived from very-long-chain n-3 PUFAs of 28 or more carbons has been identified (32, 33) and lends to the possibility that THA-derived lipid mediators and other bioactive metabolites have the potential to mediate some of the effects of DHA observed following DHA administration.

The direct plasma THA turnover (THA from plasma unesterified THA) was slower than the direct turnover of plasma DHA (Table 1). In light of the previous paragraph’s discussion, this is somewhat of a surprising finding. In order for plasma THA levels to remain low and plasma DHA to remain high, an opposite effect would have been expected. One explanation might be the tracer/tracee ratios (labeled tracer/unesterified PUFA) of the ²H₅-THA tracer that is approximately three times higher than the ¹³C₂₂-DHA tracer, meaning that 33% of all the unesterified THA present throughout the infusion is the ²H₅-THA. This may again support an “obligation” for THA conversion to DHA, as the higher than normal tracer/tracee ratio has potentially flooded the pathway to create a bottleneck at THA, which, when calculating turnover, would result in artificially slower values. This may necessitate lower infusion levels of ²H₅-THA for future studies while keeping infusate levels high enough so as to remain detectable by mass spectrometry.

In conclusion, we have shown for the first time in vivo that DHA is both a product and a precursor to THA (and vice versa) and determined daily synthesis-secretion rates for both n-3 PUFAs. Furthermore, although synthesis secretion of DHA from THA is significantly higher than THA from DHA, additional studies feeding DHA would be expected to yield more similar rates between the directions. In addition, the recent application of these infusion models to humans (24), the evidence from human cell lines suggesting the bypassing of the Sprecher pathway (4, 5), and the potentially very low ²H₅-THA tracer requirements make the implementation of human THA infusions an exciting future step. In particular, the comparison of DHA synthesis rates between individuals with peroxisomal disorders, such as Zellweger’s syndrome, would lead to an even more comprehensive understanding of such conditions.