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Keywords:

  • brain;
  • deprivation;
  • docosahexaenoic;
  • n-3 polyunsaturated fatty acids;
  • phospholipids;
  • rat

Abstract

  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Materials
  5. Animals
  6. n-3 PUFA adequate and deficient diets
  7. Intra-cerebroventricular (i.c.v.) injection of [4,5-3H]DHA
  8. Collection of brains
  9. Isolation of brain and plasma lipids
  10. Quantitation of phospholipid and fatty acid concentrations and radioactivity
  11. Fatty acid phenacyl ester preparation and HPLC analysis
  12. Calculations and statistics
  13. Results
  14. Discussion
  15. References

Male rat pups (21 days old) were placed on a diet deficient in n-3 polyunsaturated fatty acids (PUFAs) or on an n-3 PUFA adequate diet containing α-linolenic acid (α-LNA; 18 : 3n-3). After 15 weeks on a diet, [4,5-3H]docosahexaenoic acid (DHA; 22 : 6n-3) was injected into the right lateral cerebral ventricle, and the rats were killed at fixed times over a period of 60 days. Compared with the adequate diet, 15 weeks of n-3 PUFA deprivation reduced plasma DHA by 89% and brain DHA by 37%; these DHA concentrations did not change thereafter. In the n-3 PUFA adequate rats, DHA loss half-lives, calculated by plotting log10 (DHA radioactivity) against time after tracer injection, equaled 33 days in total brain phospholipid, 23 days in phosphatidylcholine, 32 days in phosphatidylethanolamine, 24 days in phosphatidylinositol and 58 days in phosphatidylserine; all had a decay slope significantly greater than 0 (p < 0.05). In the n-3 PUFA deprived rats, these half-lives were prolonged twofold or greater, and calculated rates of DHA loss from brain, Jout, were reduced. Mechanisms must exist in the adult rat brain to minimize DHA metabolic loss, and to do so even more effectively in the face of reduced n-3 PUFA availability for only 15 weeks.

Abbreviations used
AA

arachidonic acid

α-LNA

α-linolenic acid

DHA

docosahexaenoic acid

DPA

docosapentaenoic acid

FAME

fatty acid methyl ester

FAPE

fatty acid phenacyl ester

GC

gas chromatography

i.c.v.

intracerebroventricular

iPLA2

Ca2+-independent PLA2

LA

linoleic acid

PLA2

phospholipase A2

PUFA

polyunsaturated fatty acid

PC

phosphatidylcholine

PE

phosphatidylethanolamine

PI

phosphatidylinositol

PL

phospholipid

PS

phosphatidylserine

PPAR

peroxisome proliferator-activated receptors

TLC

thin layer chromatography

TNS

6-p-toluidine-2-naphthalene sulfonic acid

n-3 Polyunsaturated fatty acids (PUFAs) are dietary essential and are critical for the development and function of mammalian brain and retina. Rats, mice, piglets and monkeys show impaired brain function when deprived of n-3 PUFAs (Neuringer et al. 1986; Bourre et al. 1989; Greiner et al. 1999; Innis 2000a; Catalan et al. 2002; Champoux et al. 2002), and dietary docosahexaenoic acid (DHA; 22 : 6n-3) supplementation is reported to enhance vision and cognition in pre-term and full-term human infants (Heird et al. 1997; Carlson and Neuringer 1999; Birch et al. 2000; Makrides et al. 2000; Neuringer 2000; SanGiovanni et al. 2000; O'Connor et al. 2001; Auestad et al. 2003). Additionally, low plasma DHA levels have been associated with Alzheimer's disease and depression, whereas dietary supplementation with n-3 PUFAs may be beneficial in these diseases (Conquer et al. 2000; Mischoulon and Fava 2000; Marangell et al. 2003; Otto et al. 2003; Tully et al. 2003). Finally, plasma DHA concentrations have been reported to be decreased in elderly cohorts, and some age-related cognitive deficits have been reported to be ameliorated by n-3 PUFA supplementation (McGahon et al. 1999; Suzuki et al. 2001).

DHA must be obtained directly from the diet or be synthesized from its dietary available precursor, α-linolenic acid (α-LNA; 18 : 3n-3), by desaturation, elongation and peroxisomal β-oxidative reactions (Sprecher 2000). DHA is enriched in phospholipids of the mammalian brain and retina (Innis 2000b; Uauy et al. 2000). In these organs, it is largely esterified in the stereospecifically numbered-2 (sn-2) position of phospholipids, from where it can be released by a phospholipase A2 (PLA2). After release in brain, the unesterified DHA is reincorporated into the phospholipids (Lands and Crawford 1976; Purdon and Rapoport 1998; Farooqui et al. 2000; Rapoport 2003) or is lost by conversion to docosanoids (Kim et al. 1990; Serhan et al. 2002; Hong et al. 2003) and by β-oxidation or peroxidation (Yavin et al. 2002).

In the rat retina, the half-life for loss of DHA from phospholipids has been determined by injecting radiolabeled [4,5-3H]DHA into the vitreous humor, then measuring phospholipid radioactivity due to [4,5-3H]DHA at various times during the following 60 days. Using this method, the ‘loss’ half-life of DHA in retinal phospholipids was estimated as 19 days (Stinson et al. 1991). Furthermore, in rats subjected to n-3 PUFA deprivation following weaning, the loss half-life was prolonged to at least 1000 days. In contrast, loss half-lives of DHA have not been measured by intra-tissue injection in the mammalian brain, either under normal conditions or in response to n-3 PUFA dietary deprivation.

In view of the relevance of DHA to brain function and structure, we thought it important to determine DHA loss half-lives in brain phospholipids of n-3 PUFA dietary adequate and deprived rats. To do this, we followed much of the protocol that was used to estimate these values in the rat retina (Stinson et al. 1991). We subjected 21-day-old rat pups to 15 weeks of an n-3 PUFA adequate or deficient diet, then injected [4,5-3H]DHA into the right cerebral ventricle. Radioactivity due to [4,5-3H]DHA was determined in individual brain phospholipids at fixed times over a period of 60 days. From plots of log10 (DHA radioactivity) against time, we calculated DHA loss half-lives (T1/2) in individual brain phospholipids, as well as actual rates of DHA loss (Jout) (Rapoport et al. 2001). Part of this work has been presented in abstract form (DeMar et al. 2003, 2004a).

Materials

  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Materials
  5. Animals
  6. n-3 PUFA adequate and deficient diets
  7. Intra-cerebroventricular (i.c.v.) injection of [4,5-3H]DHA
  8. Collection of brains
  9. Isolation of brain and plasma lipids
  10. Quantitation of phospholipid and fatty acid concentrations and radioactivity
  11. Fatty acid phenacyl ester preparation and HPLC analysis
  12. Calculations and statistics
  13. Results
  14. Discussion
  15. References

4,7,10,13,16,19 [4,5-3H]Docosahexaenoic acid ([4,5-3H]DHA) in 100% ethanol, specific activity = 50 Ci/mm, was purchased from Perkin Elmer Life Sciences, NEN Life Science Products (Boston, MA, USA). High performance liquid chromatography (HPLC) with continuous scintillation counting was used to verify radioactive purity as > 98%. Sodium pentobarbital (50 mg/mL), 0.9% saline (sterile, 0.9% benzyl alcohol) and 1% lidocaine–HCl solutions were from Abbott Laboratories (North Chicago, IL, USA). HEPES [4-(2-hydroxymethyl)-1-piperazine ethane sulfonic acid], fatty acid-free bovine serum albumin, atropine methylbromide and neomycin sulfate were from Sigma-Aldrich (St Louis, MO, USA). Di-heptadecanoate phosphatidylcholine (di-17 : 0 PC) was purchased from Avanti Polar-Lipids (Alabaster, AL, USA). Thin layer chromatography (TLC) standards for mixed acyl chain phospholipids, phosphatidylcholine (PC), phosphatidylethanolamine (PE), phosphatidylserine (PS) and phosphatidylinositol (PI) were purchased from Sigma-Aldrich. Fatty acid methyl esters and free fatty acids for GC and HPLC standards were purchased from NuChek Prep (Elysian, MN, USA). 6-p-Toluidine-2-naphthalene sulfonic acid (TNS) was from Acros Organics (Fair Lawn, NJ, USA). Bromoacetophenone and triethylamine were from Fluka Chemicals (Buchs, Switzerland)/Sigma-Aldrich. All solvents were HPLC grade and from Fisher Scientific (Fair Lawn, NJ, USA) or EMD Chemicals (Gibbstown, NJ, USA). Other chemicals, unless noted, were from Fisher Scientific.

Animals

  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Materials
  5. Animals
  6. n-3 PUFA adequate and deficient diets
  7. Intra-cerebroventricular (i.c.v.) injection of [4,5-3H]DHA
  8. Collection of brains
  9. Isolation of brain and plasma lipids
  10. Quantitation of phospholipid and fatty acid concentrations and radioactivity
  11. Fatty acid phenacyl ester preparation and HPLC analysis
  12. Calculations and statistics
  13. Results
  14. Discussion
  15. References

The protocol was approved by the Animal Care and Use Committee of the National Institute on Child Health and Human Development (NICHD). Eighteen-day-old male Long-Evans rat pups and their nursing mother were purchased from Charles River Laboratories (Portage, MI, USA). The rats were housed with strictly regulated temperature (24°C) under a 12 h light/12 h dark cycle. The pups were allowed to nurse for 3 days. When 21 days old, they were removed from the mother and fed ad libitum an n-3 PUFA adequate or deficient diet, as described below. They were maintained on their diet throughout the remainder of experiment.

n-3 PUFA adequate and deficient diets

  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Materials
  5. Animals
  6. n-3 PUFA adequate and deficient diets
  7. Intra-cerebroventricular (i.c.v.) injection of [4,5-3H]DHA
  8. Collection of brains
  9. Isolation of brain and plasma lipids
  10. Quantitation of phospholipid and fatty acid concentrations and radioactivity
  11. Fatty acid phenacyl ester preparation and HPLC analysis
  12. Calculations and statistics
  13. Results
  14. Discussion
  15. References

Rodent diets (prepared by Dyets, Inc., Bethlehem, PA, USA) were designed around a standard AIN-93 formulation, with carbohydrate, protein, fat, fiber, salt and vitamin/essential amino acid content set at 60, 20, and 10, 5, 3.5, 1.5% (by weight), respectively (Reeves et al. 1993; Moriguchi et al. 2001). Ingredients and fatty acid content of n-3 PUFA adequate and deficient diets are given in Tables 1 and 2, respectively. Dietary fat came from adding select amounts of hydrogenated coconut, safflower or flaxseed oils. Hydrogenated coconut oil (approximately 6%) was added to both diets as a saturated fatty acid base. Both diets received safflower oil (approximately 3%) to provide linoleic acid (LA; 18 : 2n-6) as about 26% of total fatty acids. The safflower oil was very low in α-LNA (< 0.1%) and did not contain other n-3 PUFAs. Flaxseed oil (approximately 1%) was added to only the n-3 PUFA adequate diet to provide α-LNA at about 4% of total fatty acids. No other n-3 PUFA was found in the adequate diet. LA and α-LNA were set at 6% and 1% of total caloric intake (3935 kcal/kg), giving a ratio of 6 : 1 in the n-3 PUFA adequate diet. Intake and ratio of LA and α-LNA followed the highest recommended levels, to produce optimal arachidonic acid (AA) and DHA accretion in the brain (Bourre et al. 1989; Van Aerde and Clandinin 1993).

Table 1.  General composition (% by wt.) of n-3 PUFA adequate and deficient diets
ComponentWeight % (g /100 g diet)
n-3 Adequaten-3 Deficient
  1. a THBQ, t-butylhydroquinone.

Protein:2020
Casein2020
Carbohydrate:6060
Dextrose2020
Cornstarch1515
Maltodextrin1515
Sucrose1010
Fat:1010
Hydrogenated coconut oil6.06.6
Safflower oil3.23.4
Flaxseed oil0.8––
Additives:1010
Cellulose5.05.0
Salts3.53.5
Vitamins1.01.0
Choline chloride0.250.25
L-Cystine0.250.25
THBQa0.0020.002
Table 2.  Fatty acid composition and caloric contribution of n-3 PUFA adequate and deficient diets
Fat componentn-3 Adequaten-3 Deficient
% FatKcal/kg% Energya% FatKcal/kg% Energya
  1. Dietary PUFAs 20 : 4n-6, 20 : 5n-3, 22 : 5 n-3, and 22 : 6 n-3 were not detected (< 0.001% of total fat). aEnergy content of each fat calculated from total caloric content of each diet (3935 kcal/kg), as provided by protein (20%), carbohydrate (60%), fat (10%), and additives (10%).

Saturated63.957514.669.562615.9
Monounsaturated5.3481.24.1370.9
Linoleate (18:2n-6)26.42386.026.42386.0
Linolenate (18:3n-3)4.4401.00.040.360.009
(18:2n-6)/(18:3n-3)666660660660

Intra-cerebroventricular (i.c.v.) injection of [4,5-3H]DHA

  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Materials
  5. Animals
  6. n-3 PUFA adequate and deficient diets
  7. Intra-cerebroventricular (i.c.v.) injection of [4,5-3H]DHA
  8. Collection of brains
  9. Isolation of brain and plasma lipids
  10. Quantitation of phospholipid and fatty acid concentrations and radioactivity
  11. Fatty acid phenacyl ester preparation and HPLC analysis
  12. Calculations and statistics
  13. Results
  14. Discussion
  15. References

Rats were raised for 15 weeks (21–126 days old) on an n-3 PUFA adequate or deficient diet (n = 21). Tail blood was collected at 0, 5, 10 and 15 weeks to monitor DHA status of the two groups. Plasma was separated from blood and frozen at −80°C. At 15 weeks, animals were anesthetized by sodium pentobarbital (75 mg/kg, i.p.), and were given atropine methylbromide (5 mg/kg, i.p.; 5 mg/mL in 0.9% saline) to assist respiration. The head was placed in a stereotaxic instrument (Model 900, David Kopf Instruments; Tujunga, CA, USA) and the skull exposed. An injection needle was inserted into the right lateral cerebral ventricle (4 mm ventral to the dura) via a hole that was drilled in the cranium at 1 mm posterior and 1.5 mm lateral to the bregma (Noble et al. 1967; Gatti et al. 1986). An injection (5 µL total volume, 0.17 µL/min) was made of 40 µCi of [4,5-3H]DHA in 5 mm HEPES buffer (pH 7.4), 50 mg/mL fatty acid-free bovine serum albumin (BSA) (Gatti et al. 1986), after which the hole was sealed with cranioplastic cement and cyanoacrylate gel (Plastics one; Roanoke, VA, USA). The wound was flushed with 1% neomycin sulfate in 0.9% saline (w/v) and closed with surgical staples. For pain control, 1% lidocaine–HCl solution was applied. Subcutaneous injection of 10 mL 0.9% saline was given to prevent dehydration. Animals were allowed to recover and then were returned to their respective n-3 PUFA adequate or deficient diets.

Collection of brains

  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Materials
  5. Animals
  6. n-3 PUFA adequate and deficient diets
  7. Intra-cerebroventricular (i.c.v.) injection of [4,5-3H]DHA
  8. Collection of brains
  9. Isolation of brain and plasma lipids
  10. Quantitation of phospholipid and fatty acid concentrations and radioactivity
  11. Fatty acid phenacyl ester preparation and HPLC analysis
  12. Calculations and statistics
  13. Results
  14. Discussion
  15. References

At 2, 6, 12, 24, 36, 48 and 60 days post-i.c.v. injection of [4,5-3H]DHA, n-3 PUFA adequate and deprived rats (n = 3) were killed by an overdose of sodium pentobarbital (150 mg/kg, i.p.). To rapidly stop brain metabolism, the head was immediately subjected to focused-beam microwave irradiation (5.5 kW, 4.0 s) using an industrial microwave generator (Model S6F, Cober Electronics; Stamford, CT, USA). Brains were removed and stored at −80°C.

Additionally, 2 or 6 days following the i.c.v. injection of [4,5-3H]DHA, brains of other n-3 PUFA adequate rats were removed, frozen, and cut in coronal sections on a cryostat at − 20°C. The sections were exposed to [3H]Hyperfilm (Amersham, Arlington Heights, IL, USA) for 15–18 weeks, then developed following the manufacturer's instructions. The films were examined and compared with an atlas of the rat brain to identify specific regions (Paxinos and Watson 1987).

Isolation of brain and plasma lipids

  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Materials
  5. Animals
  6. n-3 PUFA adequate and deficient diets
  7. Intra-cerebroventricular (i.c.v.) injection of [4,5-3H]DHA
  8. Collection of brains
  9. Isolation of brain and plasma lipids
  10. Quantitation of phospholipid and fatty acid concentrations and radioactivity
  11. Fatty acid phenacyl ester preparation and HPLC analysis
  12. Calculations and statistics
  13. Results
  14. Discussion
  15. References

Total lipids from whole brain and plasma samples (tail blood) were extracted using a partition system of chloroform : methanol : 0.5 m KCl (2 : 1 : 0.75, v/v/v) (Folch et al. 1957). Di-17 : 0 PC was added as an internal standard to plasma before extraction. Brain total lipid extracts were separated into phospholipid classes (PC, PE, PI and PS) by TLC on Silica gel 60 plates (EM Separation Technologies; Gibbstown, NJ, USA) using a solvent system of chloroform : methanol : water : glacial acetic acid (60 : 50 : 4 : 1, v/v/v/v) (Chang et al. 1999). TLC plates were sprayed with 0.03% TNS in 50 mm Tris buffer (pH 7.4) (w/v) and lipid bands were visualized under UV light. The positions of brain PC, PE, PI and PS bands were identified using phospholipid standards run on the TLC plates.

Quantitation of phospholipid and fatty acid concentrations and radioactivity

  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Materials
  5. Animals
  6. n-3 PUFA adequate and deficient diets
  7. Intra-cerebroventricular (i.c.v.) injection of [4,5-3H]DHA
  8. Collection of brains
  9. Isolation of brain and plasma lipids
  10. Quantitation of phospholipid and fatty acid concentrations and radioactivity
  11. Fatty acid phenacyl ester preparation and HPLC analysis
  12. Calculations and statistics
  13. Results
  14. Discussion
  15. References

Brain phospholipid bands were scraped from TLC plates and subjected to liquid scintillation counting. Radioactivities (dpm) were adjusted for counting efficiency and converted to curies (Ci). Concentrations of brain phospholipid classes were determined by assaying the lipid phosphorus content of TLC scrapes (Rouser et al. 1970). To determine fatty acid concentrations in each phospholipid, TLC scrapes containing brain phospholipids were converted to fatty acid methyl esters (FAMEs) using 1% H2SO4 in methanol (v/v) (Makrides et al. 1994). Prior to methylation, di-17 : 0 PC was added as an internal standard to brain phospholipids. FAMEs were separated on a 30 m × 0.25 mm i.d. capillary column (SP-2330; Supelco, Bellefonte, PA, USA) using a GC with flame ionization detector (Model 6890 N; Agilent Technologies, Palo Alto, CA, USA). Runs were initiated at 80°C, with a temperature gradient to 160°C (10°C/min) and 230°C (3°C/min) in 31 min, and held at 230°C for 10 min. Peaks were identified by retention times of FAME standards. Fatty acid concentrations (µmol/g wet wt brain) were calculated by proportional comparison of GC peak areas with the area of the 17 : 0 internal standard. The fatty acid concentrations in total plasma lipid extracts were determined in a similar way.

Fatty acid phenacyl ester preparation and HPLC analysis

  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Materials
  5. Animals
  6. n-3 PUFA adequate and deficient diets
  7. Intra-cerebroventricular (i.c.v.) injection of [4,5-3H]DHA
  8. Collection of brains
  9. Isolation of brain and plasma lipids
  10. Quantitation of phospholipid and fatty acid concentrations and radioactivity
  11. Fatty acid phenacyl ester preparation and HPLC analysis
  12. Calculations and statistics
  13. Results
  14. Discussion
  15. References

Total lipid extracts obtained from brains of n-3 PUFA adequate and deprived rats (n = 3), killed at 2, 36 or 60 days post-injection of [4,5-3H]DHA, were saponified with 2% KOH/EtOH (w/v) at 100°C for 45 min, then acidified with HCl and extracted with hexane. The fatty acid extracts were converted to fatty acid phenacyl esters (FAPEs) by reacting with bromoacetophenone and triethylamine (Chen and Anderson 1992). FAPEs were separated on HPLC (Beckman, Fullerton, CA, USA) using a 25 cm × 4.6 mm i.d., C18 reverse phase column (Luna; Phenomenex, Torrence, CA, USA). Elution (2 mL/min) of FAPE was by a linear gradient of acetonitrile/water, initiated at 80 : 20 (v/v), increased to 92 : 8 (v/v) in 45 min and held at 92 : 8 (v/v) for 10 min. FAPE elution was monitored at 242 nm on a UV/VIS detector (Gilson, Middleton, WI, USA). Radioactivity profiles were obtained with an on-line flow scintillation counter (β-Ram, model 2B; IN/US Systems, Tampa, FL, USA). Peaks were identified from retention times of FAPE prepared for fatty acid standards and [4,5-3H]DHA.

Calculations and statistics

  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Materials
  5. Animals
  6. n-3 PUFA adequate and deficient diets
  7. Intra-cerebroventricular (i.c.v.) injection of [4,5-3H]DHA
  8. Collection of brains
  9. Isolation of brain and plasma lipids
  10. Quantitation of phospholipid and fatty acid concentrations and radioactivity
  11. Fatty acid phenacyl ester preparation and HPLC analysis
  12. Calculations and statistics
  13. Results
  14. Discussion
  15. References

Data were expressed as means ± SD. Log10 (radioactivity in total or individual brain phospholipid, nCi/g brain) was plotted against time post-i.c.v. injection of [4,5-3H]DHA, and the data were fit by linear regression to provide slopes in units per days. An anova was used to determine the standard error of the slope and whether it was significantly different from zero. The anova was also used to determine whether the slope in each phospholipid differed significantly between n-3 PUFA deprived and adequate rats. Statistical significance was taken as p ≤ 0.05.

Loss half-lives (days) of [4,5-3H]DHA were calculated from the measured slopes in total and individual phospholipids by the following equation (Stinson et al. 1991):

  • image(1)

Results

  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Materials
  5. Animals
  6. n-3 PUFA adequate and deficient diets
  7. Intra-cerebroventricular (i.c.v.) injection of [4,5-3H]DHA
  8. Collection of brains
  9. Isolation of brain and plasma lipids
  10. Quantitation of phospholipid and fatty acid concentrations and radioactivity
  11. Fatty acid phenacyl ester preparation and HPLC analysis
  12. Calculations and statistics
  13. Results
  14. Discussion
  15. References

Rats fed an n-3 PUFA adequate or deficient diet for 15 weeks, starting at 21 days of age, did not show a significant difference in daily weight gain, 5.6 ± 2.6 (in adequate) vs. 5.8 ± 2.8 g/day (in deprived), respectively, nor in final body weight, 660 ± 76 vs. 676 ± 72 g, respectively (n = 21). At 2–60 days post-i.c.v. [4,5-3H]DHA, their brain weights also did not differ significantly, equaling 1.83 ± 0.17 vs. 1.84 ± 0.17 g, respectively.

The total lipid DHA concentration in plasma of the n-3 PUFA adequate rats did not change during the initial 15 weeks on the diet (Fig. 1). In contrast, n-3 PUFA deprivation resulted in an 87% reduction in the plasma DHA concentration (p < 0.001) by 5 weeks, but no significant change thereafter (Fig. 1). Compared with the n-3 PUFA adequate diet, 15 weeks of n-3 PUFA deprivation reduced the plasma concentration of α-LNA by 95% (p < 0.001), while elevating by 93% (p < 0.001) the esterified concentration of the AA-elongation product, docosapentaenoic acid (DPA; 22 : 5n-6) (Table 3). Plasma LA and AA concentrations, however, were not significantly changed.

image

Figure 1. DHA concentration in plasma total lipids during 15 weeks of feeding n-3 PUFA adequate (•) or deprived (▪) diets. Data are means ± SD (n = 8).

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Table 3.   Fatty acid concentrations in plasma total lipids from n-3 PUFA adequate and deprived rats, after 0, 5, 10 and 15 weeks on diets
Esterified Fatty acidConcentration, µmol/ml plasma
0 weeks5 weeks10 weeks15 weeks
Adq.Dep.Adq.Dep.Adq.Dep.Adq.Dep.
  1. Data are means ± SD (n = 8). Fatty acids: 16 : 0, palmitic; 16 : 1n-9, palmitoleic; 18 : 0, stearic; 18 : 1n-9, oleic; 18 : 2n-6, linoleic, 18 : 3n-3, α-linolenic; 20 : 4n-6, arachidonic; 22 : 5n-6, docosapentaenoic; 22 : 6n-3, docosahexaenoic. ap < 0.05, bp < 0.01, cp < 0.001; significant difference between n-3 PUFA deprived (Dep.) and n-3 PUFA adequate (Adq.) means.

16:01.8 ± 0.61.7 ± 0.51.5 ± 0.31.4 ± 0.31.9 ± 0.41.5 ± 0.3a1.2 ± 0.41.1 ± 0.4
16:1n-90.068 ± 0.0280.056 ± 0.0180.21 ± 0.070.15 ± 0.060.28 ± 0.080.18 ± 0.07a0.16 ± 0.070.13 ± 0.07
18:01.0 ± 0.41.1 ± 0.41.3 ± 0.31.2 ± 0.11.4 ± 0.31.2 ± 0.1a1.1 ± 0.31.1 ± 0.3
18:1n-90.66 ± 0.310.55 ± 0.170.77 ± 0 .180.69 ± 0.160.99 ± 0.200.70 ± 0.26a0.52 ± 0.200.42 ± 0.19
18:2n-61.9 ± 0.72.0 ± 0.81.2 ± 0.21.3 ± 0.21.3 ± 0.31.1 ± 0.20.89 ± 0.300.80 ± 0.26
18:3n-30.036 ± 0.0110.032 ± 0.0080.036 ± 0.0090.003 ± 0.002c0.041 ± 0.0280.001 ± 0.001b0.040 ± 0.0140.002 ± 0.001c
20:4n-61.3 ± 0.61.2 ± 0.51.9 ± 0.61.9 ± 0.32.1 ± 0.42.0 ± 0.31.8 ± 0.41.9 ± 0.1
22:5n-60.012 ± 0.0030.012 ± 0.0040.015 ± 0.0040.17 ± 0.07c0.012 ± 0.0040.19 ± 0.06c0.012 ± 0.0060.18 ± 0.05c
22:6n-30.26 ± 0.120.28 ± 0.140.23 ± 0.050.029 ± 0.007c0.24 ± 0.060.026 ± 0.006c0.22 ± 0.070.023 ± 0.006c
Total7.0 ± 2.56.9 ± 2.37.2 ± 1.36.8 ± 0.98.3 ± 1.36.9 ± 1.16.0 ± 1.55.7 ± 1.5
22:5/22:60.052 ± 0.0120.048 ± 0.0130.068 ± 0.0075.6 ± 1.1c0.051 ± 0.0147.5 ± 0.6c0.053 ± 0.0167.9 ± 0.6c
n-6/n-312.5 ± 0.912.4 ± 1.714.1 ± 1.5118 ± 16c14.4 ± 1.5136 ± 14c12.4 ± 1.2127 ± 8c

On a percentage basis, less marked changes occurred in esterified DHA within brain phospholipids than in plasma total lipids. Esterified DHA in brain had fallen by 27–43% after at least 15 weeks of n-3 PUFA deprivation (Fig. 2). The brain concentration of DHA did not significantly change after the 15 weeks. The decline in brain DHA was unaccompanied by an alteration in the brain concentration of PC, PE, PI or PS (Table 4).

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Figure 2. DHA concentration in brain total phospholipids of n-3 PUFA adequate (•) and deprived (▪) rats, determined 2, 6, 12, 24, 36, 48 and 60 days post-i.c.v. injection of [4,5-3H]DHA. Data are means ± SD (n = 3).

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Table 4.  Concentrations of brain phospholipids from n-3 PUFA adequate and deprived rats, summed from 36, 48, and 60 days post-intracerebroventricular injection of [4,5-3H] DHA
Concentration, µ mol/g brain
PCPEPIPS
Adq.Dep.Adq.Dep.Adq.Dep.Adq.Dep.
  1. Data are means ± SD (n = 9). PC, phosphatidylcholine; PE, phosphatidylethanolamine; PS, phosphatidylserine; PI, phosphatidylinositol. There were no significant differences detected between n-3 PUFA deprived (Dep.) and n-3 PUFA adequate (Adq.) means.

24.5 ± 1.223.8 ± 1.525.1 ± 1.924.6 ± 0.92.7 ± 0.32.5 ± 0.37.8 ± 0.67.7 ± 0.4

Table 5 summarizes esterified fatty acid concentrations in brain PC, PE, PI, PS and total phospholipids of the two diet groups, taken as a whole over the post-injection period. In neither group was α-LNA detected. All brain phospholipid classes in the n-3 PUFA deprived compared with the adequate rats had significantly less DHA. The DHA concentration was reduced by 37% in total phospholipids (p < 0.001), with the highest fractional depletion (49%) in PS (p < 0.001). PC, PE and PI showed 39%, 33% and 20% lower levels, respectively (p < 0.05). In contrast, the concentration of DPAn-6 was increased by 87–95% (p < 0.001) in these phospholipids. LA, the dietary-essential precursor of AA, was decreased in PC and PS by 23% and 39% (p < 0.001), respectively, but was not significantly changed in PE or PI. AA itself was reduced in PS by 18% (p < 0.05), but was not altered significantly in PC, PE or PI. There was a small but significant 10% decrease of oleic acid (18 : 1n-9) in PE (p < 0.05), and 8–44% reductions of certain saturated and mono-unsaturated fatty acids in PC and PS (p < 0.05).

Table 5.  Esterified fatty acid concentrations in brain phospholipid classes from n-3 PUFA adequate and deprived rats, measured 2–60 days post intra-cerebroventricular of [4,5-3H]DHA
Esterified Fatty acidConcentration, µmol/g brain
PCPEPIPSTotal PL
Adq.Dep.Adq.Dep.Adq.Dep.Adq.Dep.Adq.Dep.
  1. Data are means ± SD (n = 21). Fatty acid and phospholipid abbreviations (see Fig. legend 3 and Table legend 4, respectively). ND, not detected. ap < 0.05, bp < 0.01, cp < 0.001; significant difference between n-3 PUFA deprived (Dep.) and n-3 PUFA adequate (Adq.) means.

16:020.6 ± 1.819.0 ± 2.5a2.6 ± 0.22.7 ± 0.30.38 ± 0.080.38 ± 0.100.26 ± 0.060.22 ± 0.1123.8 ± 1.822.3 ± 2.7a
16:1n-90.24 ± 0.030.22 ± 0.03b0.13 ± 0.020.13 ± 0.040.010 ± 0.0040.008 ± 0.0030.015 ± 0.0120.008 ± 0.004a0.40 ± 0.040.37 ± 0.06a
18:06.9 ± 0.46.5 ± 0.5b7.7 ± 0.97.8 ± 0.81.4 ± 0.31.6 ± 0.56.1 ± 1.14.8 ± 1.9a22.1 ± 2.120.6 ± 3.2
18:1n-910.9 ± 0.610.1 ± 0.6c8.4 ± 1.27.5 ± 0.8a0.36 ± 0.140.43 ± 0.223.2 ± 0.62.4 ± 1.0b22.9 ± 1.820.5 ± 2.1c
18:2n-60.36 ± 0.020.28 ± 0.03c0.16 ± 0.050.14 ± 0.050.017 ± 0.0060.017 ± 0.0100.030 ± 0.0100.018 ± 0.010c0.57 ± 0.050.46 ± 0.07c
18:3n-3NDNDNDNDNDNDNDNDNDND
20:4n-62.4 ± 0.22.5 ± 0.35.1 ± 0.75.3 ± 0.61.3 ± 0.31.5 ± 0.50.54 ± 0.090.44 ± 0.18a9.4 ± 1.19.8 ± 1.5
22:5n-60.036 ± 0.0060.74 ± 0.26c0.13 ± 0.042.6 ± 0.9c0.010 ± 0.0060.059 ± 0.032c0.077 ± 0.0281.0 ± 0.67c0.25 ± 0.064.4 ± 1.8c
22:6n-31.7 ± 0.31.1 ± 0.2c7.3 ± 1.64.9 ± 0.7c0.15 ± 0.030.12 ± 0.05a2.8 ± 0.71.5 ± 0.69c12.0 ± 2.47.6 ± 1.5c
22:5/22:60.020 ± 0.0020.67 ± 0.17c0.017 ± 0.0030.52 ± 0.13c0.073 ± 0.0520.48 ± 0.14c0.026 ± 0.0040.66 ± 0.17c0.021 ± 0.0030.57 ± 0.15c
n-6/n-31.6 ± 0.153.2 ± 0.3c0.75 ± 0.061.6 ± 0.2c9.3 ± 0.714.2 ± 1.8c0.23 ± 0.030.98 ± 0.17c0.86 ± 0.081.9 ± 0.2c

Two and 6 days after injecting [4,5-3H]DHA into the right lateral cerebral ventricle of additional n-3 PUFA adequate rats, brains were removed, frozen, and subjected to quantitative [3H]-autoradiography. At both times, the right hemisphere was seen to be widely labeled, with dense labeling along the lateral ventricle wall and deep into the striatum (data not shown). The left hemisphere showed less labeling, with most near its lateral ventricle wall and striatum.

Percentage recovery of injected radioactivity in brain phospholipids at 2 days post-i.c.v. [4,5-3H]DHA injection was 22 ± 2 and 21 ± 2%/g brain in n-3 PUFA adequate and deprived animals, respectively (n = 3). In both groups, all fatty acid radioactivity was in the form of [4,5-3H]DHA. This was demonstrated by showing that brain radioactivity co-eluted on HPLC as a single peak with unlabeled DHA at 60 days (Fig. 3a) as well as at 2 and 36 days (data not shown) post-injection. There was no evidence of significant non-volatile, non-long-chain fatty acid oxidation products, since less than 0.5% of radioactivity recovered from brain was in the water-soluble fraction (data not shown).

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Figure 3. (a) UV absorbance (242 nm) trace vs. time (min) for HPLC of fatty acid phenacyl esters (FAPE) in brain total lipids from an n-3 PUFA adequate rat, 60 days post-i.c.v. [4,5-3H]DHA. 16 : 0, palmitic; 18 : 0, stearic; 18 : 1n-9, oleic; 20 : 4n-6, arachidonic; and DHA, docosahexaenoic acids (22 : 6n-3). (b) Radioactivity trace ([3H]-dpm × 103) vs. time (min) for HPLC, shown in (a). (c) HPLC radioactivity trace for FAPE from brain total lipids of an n-3 PUFA deprived rat (60 days post-injection); absorbance trace not shown.

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Plots of log10 (DHA radioactivity) against days post-i.c.v. injection of [4,5-3H]DHA are presented in Fig. 4 for total phospholipid and in Fig. 5 for the four phospholipid classes. The data were fit by linear regression, from which slopes and their statistical significance were calculated. The slopes for loss of radioactivity in n-3 PUFA adequate rats differed significantly from zero in total phospholipids (p < 0.001) and in each of the four phospholipid classes (p < 0.01). In n-3 PUFA deprived rats, the disappearance rates for DHA radioactivity (slopes) were reduced significantly compared with their respective values in the n-3 PUFA adequate animals. All the slopes differed significantly from zero (p < 0.01), except that for PS.

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Figure 4. Linear regression plots of log10 (DHA radioactivity, nCi/g brain) against time following i.c.v. injection of [4,5-3H]DHA, in total brain phospholipid from n-3 PUFA adequate (left panel, •) and deprived (right panel, ▪) rats, following 15 weeks on diets. Each time point has 3 rats. The slope (S) and its standard error for each regression line are shown. Slopes marked by an asterisk (*) differ significantly from zero (*p < 0.05, ***p < 0.001). ++p < 0.01, significantly different from slope of regression line for n-3 PUFA adequate rat brain. Insets in left hand corner of each plot show DHA radioactivity vs. time before log rhythmic (linear) transformation; data are fitted with log rhythmic regression lines.

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image

Figure 5. Linear regression plots of log10 (DHA radioactivity, nCi/g brain) against time, following i.c.v. injection of [4,5-3H]DHA, in individual brain phospholipids from n-3 PUFA adequate (left panel, •) and deprived (right panel, ▪) rats, following 15 weeks on diets. Each time point has 3 rats. The slope (S) and its standard error for each regression line are shown. **p < 0.01, ***p < 0.001; slopes differ significantly from zero (NS, not significant from zero). +p < 0.05, ++p < 0.01, +++p < 0.001; significantly different from slope of regression line for n-3 PUFA adequate rat brain.

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Loss half-lives for total phospholipids, and for each of the four phospholipid classes, were calculated using eqn 1 (see Methods) from the experimentally determined slopes. They are presented in Table 6, together with their corresponding half-lives, for total and individual brain phospholipids. In the n-3 PUFA adequate rats, the [4,5-3H]DHA loss half-life in the phospholipids, corresponding to statistically significant slopes, ranged from 23 days in PC to 58 days in PS. In the n-3 PUFA deficient rats, the loss half-life ranged from 51 days in PC to 77 days in PE, but it could not be estimated in PS because the slope, in the denominator of eqn 1, did not differ significantly from zero. The half-life for total phospholipids equaled 90 days, compared with 33 days in n-3 PUFA adequate rats. The slopes in PC, PE, PI and PS, as well as in total phospholipids, were significantly less in the n-3 PUFA deprived than adequate rats (p < 0.05); thus, the corresponding DHA loss half-lives were significantly prolonged by n-3 PUFA deprivation.

Table 6.  Loss half-lives of [4,5-3H]DHA from brain phospholipids
n-3 PUFA diet groupBrain lipidSlope, days−1 SE, days−1p valueT1/2, daysJout µmol/g brain/day
  1. a Slopes for decline of [4,5-3H]DHA radioactivity in brain phospholipids, and corresponding calculated loss half-lives (T1/2) and rates of loss of DHA (Jout). The latter were calculated by eqns 2 and 3, respectively. p-Values designate significance of difference of slope from 0. *p < 0.05, **p < 0.01, ***p < 0.001; differs significantly from slope in n-3 PUFA adequate rats. Abbreviations, see Table 4. PL, phospholipid. ND, not determined.

Adequate:Total PL−0.00920.0014< 0.0001330.257
PC−0.01330.0018< 0.0001230.054
PE−0.00950.0014< 0.0001320.160
PI−0.01260.0015< 0.0001240.004
PS−0.00520.00140.0015580.034
Deprived:Total PL−0.0033*0.00120.011900.058
PC−0.0059**0.00150.0012510.015
PE−0.0039**0.00120.0037770.044
PI−0.0058***0.0012< 0.0001520.002
PS+0.0003*0.00160.85NDND

The half-lives can be used to calculate actual rates of loss of esterified DHA from a phospholipid, Jout (µmol/g brain/day), as follows (Rapoport et al. 2001):

  • image(2)

where CDHA = unlabeled DHA concentration in a phospholipid (µmol/g brain). Insertion of the half-lives in Table 6 and the unlabeled DHA concentrations in Table 5 into this equation, for each phospholipid, gave the rates of loss in the last data column of Table 6. These values range from 0.004 µmol/g brain/day in PI to 0.16 µmol/g brain/day in PC, and equaled 0.257 µmol/g brain/day for total phospholipids. Overall, they were reduced fourfold by n-3 PUFA deprivation.

Discussion

  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Materials
  5. Animals
  6. n-3 PUFA adequate and deficient diets
  7. Intra-cerebroventricular (i.c.v.) injection of [4,5-3H]DHA
  8. Collection of brains
  9. Isolation of brain and plasma lipids
  10. Quantitation of phospholipid and fatty acid concentrations and radioactivity
  11. Fatty acid phenacyl ester preparation and HPLC analysis
  12. Calculations and statistics
  13. Results
  14. Discussion
  15. References

Marked, statistically significant changes in brain n-3 and n-6 PUFA composition and DHA kinetics were produced in 21-day-old rats subjected to dietary n-3 PUFA deprivation for 15 weeks. These changes, which had reached a new steady-state by 15 weeks, included reductions in the esterified DHA concentration in individual brain phospholipids and compensatory increases in the esterified DPAn-6 concentration. Slopes of [4,5-3H]DHA loss in total and individual brain phospholipids were significantly reduced by n-3 PUFA deprivation (p < 0.05), and corresponding loss half-lives were prolonged twofold or greater. Their prolongation corresponded to reduced rates of consumption (Jout) of the unlabeled DHA in each phospholipid. For total phospholipid, Jout was reduced fourfold. There was no significant effect of n-3 PUFA deprivation on the brain concentration of PC, PE, PI or PS, suggesting that the changes in DHA loss were not due to altered turnover rates and/or de novo synthesis rates of the phospholipid glycerol backbone alone.

Behavioral and brain structural effects of n-3 PUFA deprivation have been ascribed to an imbalance among brain concentrations of 20 : 4n-6, 22 : 5n-6 and 22 : 6n-3, or their bioactive metabolites (Contreras et al. 2001; Salem & Nieblylski, 1995). Reductions in brain DHA have been shown to modulate gene expression (Khair-El-Din et al. 1996), enzyme activity (Martin 1998), membrane channels (Poling et al. 1995; Hamano et al. 1996), and formation of prostaglandins, leukotrienes (Corey et al. 1983; Matsumoto et al. 1993) and neuroprostane (Reinboth et al. 1996). An n-3 PUFA deficient diet can also alter brain neuroreceptor densities and neurotransmission (Delion et al. 1996; Chalon et al. 1998).

In the present study, a 37% reduction in brain esterified DHA, from 12.0 µmol/g brain to 7.6 µmol/g brain, was produced by 15 weeks of n-3 PUFA deprivation in a single (F1) rat generation, compared with a > 75% reduction following n-3 PUFA deprivation in three generations (F3) of rats (Contreras et al. 2000; Catalan et al. 2002). No significant change after 15 weeks was evident in the brain PS concentration, although a PS reduction has been reported after three generations of n-3 PUFA deprivation in rats (Hamilton et al. 2000; Murthy et al. 2002). Esterified DPAn-6 was increased in our study, largely replacing the DHA that was lost. Similar but larger increments in DPAn-6, also largely replacing lost DHA, have been reported following three generations of n-PUFA deprivation (Bourre et al. 1989; Gazzah et al. 1995; Salem and Nieblylski 1995; Contreras et al. 2000, 2001; Catalan et al. 2002). As with the more extreme deprivation of three generations (Contreras et al. 2000), we found no significant total changes in the brain esterified AA concentration.

The brain growth spurt, when myelin and synapses are laid down, is finished in rats at 30 days of age (Su et al. 1996). We initiated n-3 PUFA deprivation at 21 days, when the growing brain was still accreting DHA into newly made membranes. This is likely why we obtained a significant depletion of brain DHA (37%). In contrast to this finding, 60-day-old rats that were deprived of n-3 PUFA for up to 7 months showed no significant depletion of brain DHA (Bourre et al. 1992). This lack of loss likely was due to brain incorporation of DHA released from whole body stores mainly in liver and adipose tissue, as well as to reduced turnover (Su et al. 1996). Similar considerations would apply to starting n-3 PUFA deprivation at any time after the brain growth spurt.

HPLC analysis of long-chain fatty acid radioactivity at 2–60 days following i.c.v. [4,5-3H]DHA (Fig. 3), and measurement of aqueous metabolites in extracted brain, showed that [4,5-3H]DHA metabolites, or its elongated or shortened products, were essentially absent from brain, as has been reported for the retina (Stinson et al. 1991). Retro-conversion of DHA is minor, and does not pass beyond the formation of small amounts of docosapentaenoic acid (DPA; 22 : 5n-3) and eicosapentaenoic acid (EPA; 20 : 5n-3). In studies on rats (Brossard et al. 1996) designed to address DHA retroconversion, orally administered [13C]-DHA gave maximal plasma stable-isotope concentrations (2 h post-dose) of DPA and EPA at 2% and 7% of the dose, respectively, mainly in the phospholipid fraction. DHA metabolism obviously occurred in the liver following oral administration, whereas direct intracerebral injection in our study by-passed liver metabolism. Our results suggest that brain, unlike liver, lacks enzymes required for n-3 PUFA retro-conversion. Thus, our slope method for calculating loss half-lives (eqn 1) was not contaminated by radioactivity not representative of intact [4,5-3H]DHA. The loss half-lives in n-3 PUFA adequate rats ranged from 23 to 58 days for the individual phospholipid classes, and equaled 33 days for the total phospholipid compartment (Table 6). They were prolonged twofold or greater by 15 weeks of n-3 PUFA deprivation.

The loss half-life of DHA in total phospholipid in the rat retina, determined by the slope method following injection of [4,5-3H]DHA into the vitreous humor, equals 19 days (Stinson et al. 1991). It is reasonably comparable with our value for brain (33 days) following 15 weeks of an n-3 PUFA adequate diet. Nevertheless, dissipation mechanisms involving DHA in the retina can differ from those in brain. Membranes containing DHA are continually shed from retinal photoreceptors and are digested by the retinal pigment epithelium (Young 1976); the pigment epithelium recycles the DHA from scavenged photoreceptor membranes back to the retina (Bazan et al. 1994).

It also is possible to estimate PUFA half-lives by ‘pulse labeling’ brain phospholipids at a single time point in an experiment (Robinson et al. 1992; Rapoport 2001, 2003), rather than by using the multiple time-point slope approach illustrated in Figs 4 and 5 that has been employed widely for long-chain fatty acids (Sun 1977; Sun and Su 1979; Porcellati et al. 1983; Stinson et al. 1991). The ‘pulse labeling’ approach involves the one-time intravenous infusion of radiolabeled PUFA for 5 min in awake rats, then killing the animal and microwaving and removing its brain. PUFA incorporation coefficients k* are calculated as the ratio of brain phospholipid radioactivity to integrated plasma radioactivity (input function). Multiplying k* by the unesterified plasma PUFA concentration gives the rate of unlabeled PUFA incorporation into phospholipid, Jin (µmol/g brain/unit time), from which a half-life can be determined as:

  • image(3)

Using one-time pulse labeling in awake adult rats (n-3 PUFA adequate), two studies reported that the half-life of total phospholipid equaled 77–111 days, whereas half-lives for PC, PE, PI and PS were 24–33 days, 85–159 days, 6–7 days and 347–596 days, respectively (Chang et al. 1999; Contreras et al. 2000). Comparison with Table 6 shows that the pulse-labeling and slope half-lives agree most closely for total phospholipid and PC, but that the slope half-life is longer than the pulse-labeling half-life for PI, and shorter than that for PE or PS. These differences, although not dramatic considering methodology differences, may arise in part because pulse-labeling assumes that, within the 5 min of intravenous tracer infusion, DHA from plasma replaces all the DHA that has been released from a phospholipid by PLA2, and that the phospholipid class is a rapidly exchanging homogeneous compartment. This assumption may not be applicable particularly to PE and PS, which are labeled from plasma slowly and indirectly through intermediate compartments, as shown using [1-14C]AA (DeGeorge et al. 1989). Additionally, pulse-labeling ignores slow incorporation by de novo synthesis of phospholipid (Porcellati et al. 1983; Horrocks 1985). The shorter pulse-labeling than slope-derived DHA half-life in PI may reflect the fact that the pulse labeling value is biased to rapidly turning over PI molecular species (Shetty et al. 1996).

Published pulse-labeling values for Jin for DHA are 0.15–0.19 µmol/g brain/day for total phospholipids, 0.036–0.054 µmol/g brain/day for PC, 0.090–0.11 µmol/g brain/day for PE, 0.017–0.019 µmol/g brain/day for PI and 0.004–0.006 µmol/g brain/day for PS (Chang et al. 1999; Contreras et al. 2000). These are roughly comparable with the slope-derived DHA values for Jout (Table 6). Indeed, because brain DHA cannot be synthesized de novo, and plasma α-LNA makes a negligible contribution to it (Moore et al. 1991; Innis and Dyer 2002; DeMar et al. 2004b), under the steady-state conditions following 15 weeks on the n-3 PUFA deficient or adequate diet, Jin should approximate Jout. Knowing this equivalency allows us to interpret Jin as also representing DHA metabolic loss from brain. This is particularly relevant, because regional values of Jin can be obtained with the pulse method using quantitative autoradiography in unanesthetized rats (Rapoport 2003) or positron emission tomography in humans (Giovacchini et al. 2002; Giovacchini et al. 2004).

Loss half-lives of AA from brain phospholipids, measured in rodents by the slope method, are also reported to be of the order of weeks (Sun 1977; Sun and Su 1979; Porcellati et al. 1983). Furthermore, for both AA and DHA, loss-half lives calculated by the slope method (Figs 4 and 5; Table 6) (Sun 1977; Sun and Su 1979; Stinson et al. 1991) or by pulse-labeling (DeGeorge et al. 1989, 1991; Chang et al. 1999; Contreras et al. 2000), are 20–50-fold longer than half-lives of actual rates of PUFA release and re-incorporation (acylation-deacylation recycling) within rodent brain phospholipids (Lands and Crawford 1976; Purdon and Rapoport 1998; Farooqui et al. 2000; Rapoport et al. 2001; Rapoport 2003). ‘Actual’ recycling half-lives can be determined by pulse-labeling during labeled PUFA infusion, by dividing Jin by the steady-state ratio, defined as λ, of the specific radioactivity of the relevant brain acyl-CoA relative to the specific activity of the plasma unesterified fatty acid. The ‘recycling’ half-life is much shorter than the ‘loss’ half-life because a large fraction (96–98%) of a PUFA released from the sn-2 position of a phospholipid, whether it be DHA or AA, is returned to the phospholipid via acyl-CoA synthetase and a lysophospholipid acyl-CoA transferase-mediated reaction, without being lost by metabolism (Rapoport et al. 2001; Rapoport 2003). Short ‘recycling’ half-lives are consistent with PUFA participation in active signal transduction (DeGeorge et al. 1989; Axelrod 1990; Shimizu and Wolfe 1990; Jones et al. 1997).

Our n-3 PUFA adequate rats, fed α-LNA but not DHA, had concentrations of esterified plasma and esterified brain DHA that were 0.23 ± 0.07 µmol/mL and 12.0 ± 2.4 µmol/g brain, respectively, which are comparable to DHA concentrations in rats whose diets contain adequate amounts of both DHA and α-LNA (0.32 ± 0.05 µmol/mL and 9.0 ± 0.5 µmol/g for plasma and brain, respectively) (Moriguchi et al. 2001). While our n-3 PUFA adequate rats attained normal brain DHA levels found in adult rats (Moriguchi et al. 2001; Rapoport et al. 2001), it has been reported that brain DHA is higher in neonatal rats provided DHA in their diet compared with rats fed only high amounts of α-LNA (Woods et al. 1996; Ward et al. 1998).

It is well documented that α-LNA is not efficiently converted to DHA, which is why we chose to provide α-LNA at a high dietary level in the n-3 PUFA adequate rats (1% of caloric intake). In rats, α-LNA is mainly shuttled to β-oxidation (> 75%) and is minimally used to synthesize DHA (< 5%) (Cunnane et al. 1999). In primates, dietary DHA is seven times more efficient than dietary α-LNA at providing DHA to the growing brain (Su et al. 1999). Likewise, adult humans show rather insignificant plasma-fractional conversion of α-LNA to DHA (< 0.1%), even with poor n-3 PUFA intake (Pawlosky et al. 2003). In cell culture, astrocytes have been shown to synthesize DHA from α-LNA and to secrete it to neurons (Moore 2001; Williard et al. 2001a,b), but this process appears insignificant in vivo in the adult rat brain (Moore et al. 1991; Innis and Dyer 2002; DeMar et al. 2004b).

Long baseline loss half-lives and their prolongation by 15 weeks of n-3 PUFA deprivation, in retina and brain, suggest that mechanisms exist to conserve DHA within these organs and that these mechanisms are up-regulated during deprivation. They may involve enzymes with a preference for DHA over AA, as n-3 PUFA deprivation even for three generations does not alter AA composition or AA recycling within rat brain phospholipids (Contreras et al. 2001). Candidate enzymes are acyl-CoA synthetases, lysophospholipid acyl-CoA transferases and PLA2s, which regulate acylation-deacylation recycling. There is no evidence, however, for DHA selectivity over AA in the acyl-CoA synthetase or lysophospholipid acyl-CoA transferase classes of enzymes (Laposata et al. 1985; Masuzawa et al. 1989; MacDonald and Sprecher 1991; Kang et al. 1997; Farooqui et al. 2000), but a Ca2+-independent PLA2 (iPLA2) is more selective for DHA than AA (Farooqui and Horrocks 2001; Strokin et al. 2003).

When released by a PLA2, DHA can be irreversibly lost by conversion to docosanoids (prostaglandin-‘like’ molecules) in part by cyclooxygenase 2 (COX-2) (Hong et al. 2003). Our n-3 PUFA deprived rats accumulated DPAn-6 in their brain phospholipids. Elevated concentrations of DPAn-6 could compete with DHA for the active sites of both PLA2 and COX-2, thus helping to retain DHA. The specificities of PLA2 and COX-2 have not been tested for DPAn-6, but in general, these enzymes appear to have high selectivity for PUFAs in the range of C20 to C22, with 4–6 double bonds (Diez et al. 1992; Ringbom et al. 2001; Strokin et al. 2003).

Reduced DHA loss during n-3 PUFA deprivation could also result from reduced activities of peroxisomal or mitochondrial oxidative enzymes, or reduced exposure to reactive oxygen species. Indeed, reduced oxidation of LA has been reported during n-6 PUFA deprivation (Cunnane and Anderson 1997). The transcription of genes of many oxidative enzymes is controlled by transcription factors known as PPARs (peroxisome proliferator-activated receptors), which can be activated by binding to an n-3 PUFA (Clarke et al. 1999; Price et al. 2000). In neuronal cultures supplied with α-LNA, esterified DHA was elevated and PPAR type δ and oxidative enzyme expression were up-regulated (Langelier et al. 2003). In contrast, n-3 PUFA deprivation for three generations did not change the brain expression of PPAR types α and β (Rojas et al. 2002). DHA oxidation could also be decreased through post-translational modification of oxidative enzymes whose expression levels are most responsive to changes in n-3 PUFA status, i.e. acyl-CoA oxidase, carnitine O-palmitoyltranferase and mitochondrial uncoupling protein (Clarke et al. 1999; Price et al. 2000).

In summary, the slope method showed that loss half-lives of DHA in individual phospholipids of the n-3 PUFA adequate rat brain ranged from 23 to 56 days. When rats were subjected to n-3 PUFA deprivation for 15 weeks post-weaning, these half-lives were prolonged twofold or greater, while the concentration of esterified DHA in brain phospholipids was reduced by an average of 37%. Studying this single generation rat model should be important for understanding the effects on brain of n-3 PUFA insufficiency in humans (Carlson and Neuringer 1999; Conquer et al. 2000; Harel et al. 2001).

References

  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Materials
  5. Animals
  6. n-3 PUFA adequate and deficient diets
  7. Intra-cerebroventricular (i.c.v.) injection of [4,5-3H]DHA
  8. Collection of brains
  9. Isolation of brain and plasma lipids
  10. Quantitation of phospholipid and fatty acid concentrations and radioactivity
  11. Fatty acid phenacyl ester preparation and HPLC analysis
  12. Calculations and statistics
  13. Results
  14. Discussion
  15. References
  • Auestad N., Scott D. T., Janowsky J. S. et al. (2003) Visual, cognitive, and language assessments at 39 months: a follow-up study of children fed formulas containing long-chain polyunsaturated fatty acids to 1 year of age. Pediatrics 112, 177183.
  • Axelrod J. (1990) Receptor-mediated activation of phospholipase A2 and arachidonic acid release in signal transduction. Biochem. Soc. Trans. 18, 503507.
  • Bazan N. G., Rodriguez de Turco E. B. and Gordon W. C. (1994) Docosahexaenoic acid supply to the retina and its conservation in photoreceptor cells by active retinal pigment epithelium-mediated recycling. World Rev. Nutr. Diet 75, 120123.
  • Birch E. E., Garfield S., Hoffman D. R., Uauy R. and Birch D. G. (2000) A randomized controlled trial of early dietary supply of long-chain polyunsaturated fatty acids and mental development in term infants. Dev. Med. Child. Neurol. 42, 174181.
  • Bourre J. M., Francois M., Youyou A., Dumont O., Piciotti M., Pascal G. and Durand G. (1989) The effects of dietary α-linolenic acid on the composition of nerve membranes, enzymatic activity, amplitude of electrophysiological parameters, resistance to poisons and performance of learning tasks in rats. J. Nutr. 119, 18801892.
  • Bourre J. M., Dumont O. S., Piciotti M. J., Pascal G. A. and Durand G. A. (1992) Dietary alpha-linolenic acid deficiency in adult rats for 7 months does not alter brain docosahexaenoic acid content, in contrast to liver, heart and testes. Biochim. Biophys. Acta 1124, 119122.
  • Brossard N., Croset M., Pachiaudi C., Riou J. P., Tayot J. L. and Lagarde M. (1996) Retroconversion and metabolism of [13C]22: 6n-3 in humans and rats after intake of a single dose of [13C]22: 6n-3-triacylglycerols. Am. J. Clin. Nutr. 64, 577586.
  • Carlson S. E. and Neuringer M. (1999) Polyunsaturated fatty acid status and neurodevelopment: a summary and critical analysis of the literature. Lipids 34, 171178.
  • Catalan J., Moriguchi T., Slotnick B., Murthy M., Greiner R. S. and Salem N., Jr (2002) Cognitive deficits in docosahexaenoic acid-deficient rats. Behav. Neurosci. 116, 10221031.
  • Chalon S., Delion-Vancassel S., Belzung C., Guilloteau D., Leguisquet A. M., Besnard J. C. and Durand G. (1998) Dietary fish oil affects monoaminergic neurotransmission and behavior in rats. J. Nutr. 128, 25122519.
  • Champoux M., Hibbeln J. R., Shannon C., Majchrzak S., Suomi S. J., Salem N., Jr and Higley J. D. (2002) Fatty acid formula supplementation and neuromotor development in rhesus monkey neonates. Pediatr. Res. 51, 273281.
  • Chang M. C., Bell J. M., Purdon A. D., Chikhale E. G. and Grange E. (1999) Dynamics of docosahexaenoic acid metabolism in the central nervous system: lack of effect of chronic lithium treatment. Neurochem. Res. 24, 399406.
  • Chen H. and Anderson R. E. (1992) Quantitation of phenacyl esters of retinal fatty acids by high-performance liquid chromatography. J. Chromatogr. 578, 124129.
  • Clarke S. D., Thuillier P., Baillie R. A. and Sha X. (1999) Peroxisome proliferator-activated receptors: a family of lipid-activated transcription factors. Am. J. Clin. Nutr. 70, 566571.
  • Conquer J. A., Tierney M. C., Zecevic J., Bettger W. J. and Fisher R. H. (2000) Fatty acid analysis of blood plasma of patients with Alzheimer's disease, other types of dementia, and cognitive impairment. Lipids 35, 13051312.
  • Contreras M. A., Greiner R. S., Chang M. C., Myers C. S., Salem N., Jr and Rapoport S. I. (2000) Nutritional deprivation of alpha-linolenic acid decreases but does not abolish turnover and availability of unacylated docosahexaenoic acid and docosahexaenoyl-CoA in rat brain. J. Neurochem. 75, 23922400.
  • Contreras M. A., Chang M. C., Rosenberger T. A., Greiner R. S., Myers C. S., Salem N., Jr and Rapoport S. I. (2001) Chronic nutritional deprivation of n-3 alpha-linolenic acid does not affect n-6 arachidonic acid recycling within brain phospholipids of awake rats. J. Neurochem. 79, 10901099.
  • Corey E. J., Shih C. and Cashman J. R. (1983) Docosahexaenoic acid is a strong inhibitor of prostaglandin but not leukotriene biosynthesis. Proc. Natl Acad. Sci. USA 80, 35813584.
  • Cunnane S. C. and Anderson M. J. (1997) Pure linoleate deficiency in the rat: influence on growth, accumulation of n-6 polyunsaturates, and [1–14C]linoleate oxidation. J. Lipid Res. 38, 805812.
  • Cunnane S. C., Menard C. R., Likhodii S. S., Brenna J. T. and Crawford M. A. (1999) Carbon recycling into de novo lipogenesis is a major pathway in neonatal metabolism of linoleate and alpha-linolenate. Prostaglandins Leukot. Essent. Fatty Acids 60, 387392.
  • DeGeorge J. J., Noronha J. G., Bell J. M., Robinson P. and Rapoport S. I. (1989) Intravenous injection of [1–14C]arachidonate to examine regional brain lipid metabolism in unanesthetized rats. J. Neurosci. Res. 24, 413423.
  • DeGeorge J. J., Nariai T., Yamazaki S., Williams W. M. and Rapoport S. I. (1991) Arecoline-stimulated brain incorporation of intravenously administered fatty acids in unanesthetized rats. J. Neurochem. 56, 352355.
  • Delion S., Chalon S., Guilloteau D., Besnard J. C. and Durand G. (1996) Alpha-linolenic acid dietary deficiency alters age-related changes of dopaminergic and serotoninergic neurotransmission in the rat frontal cortex. J. Neurochem. 66, 15821591.
  • DeMar J. C., Jr, Ma K., Bell J. and Rapoport S. I. (2003) Half-lives of docosahexaenoate in rat brain phospholipids are prolonged by nutritional deprivation of n-3 polyunsaturated fatty acids, in Gordon Research Conference Abstracts (36), Lipids and Molecular and Cellular Biology, Kimball Union Academy, NH.
  • DeMar J. C., Jr, Ma K., Bell J. M. and Rapoport S. I. (2004a) Half-lives of docosahexaenoic acid in rat brain phospholipids are prolonged by 15 weeks of deprivation of n-3 polyunsaturated fatty acids, in ISSFAL 6th Congress Abstracts, Lipids as Determinant of Cell Function and Health, Abstract 6-1. Brighton, England (UK).
  • DeMar J. C., Jr, Ma K., Bell J. M. and Rapoport S. I. (2004b) α-Linonenic acid does not contribute appreciably to the brain synthesis of docosahexaenoic acid in the adult rat, in ISSFAL 6th Congress Abstracts, Lipids as Determinant of Cell Function and Health, Abstract E2. Brighton, England (UK).
  • Diez E., Louis-Flamberg P., Hall R. H. and Mayer R. J. (1992) Substrate specificities and properties of human phospholipases A2 in a mixed vesicle model. J. Biol. Chem. 267, 18 34218 348.
  • Farooqui A. A. and Horrocks L. A. (2001) Plasmalogens: workhorse lipids of membranes in normal and injured neurons and glia. Neuroscientist 7, 232245.
  • Farooqui A. A., Horrocks L. A. and Farooqui T. (2000) Deacylation and reacylation of neural membrane glycerophospholipids. J. Mol. Neurosci. 14, 123135.
  • Folch J., Lees M. and Sloane Stanley G. H. (1957) A simple method for the isolation and purification of total lipides from animal tissues. J. Biol. Chem. 226, 497509.
  • Gatti C., Noremberg K., Brunetti M., Teolato S., Calderini G. and Gaiti A. (1986) Turnover of palmitic and arachidonic acids in the phospholipids from different brain areas of adult and aged rats. Neurochem. Res. 11, 241252.
  • Gazzah N., Gharib A., Croset M., Bobillier P., Lagarde M. and Sarda N. (1995) Decrease of brain phospholipid synthesis in free-moving n-3 fatty acid deficient rats. J. Neurochem. 64, 908918.
  • Giovacchini G., Chang M. C., Channing M. A. et al. (2002) Brain incorporation of [11C]arachidonic acid in young healthy humans measured with positron emission tomography. J. Cereb. Blood Flow Metab. 22, 14531462.
  • Giovacchini G., Lerner A., Toczek M. T., Fraser C., Ma K., DeMar J. C., Herscovitch P., Eckelman W. C., Rapoport S. I. and Carson R. E. (2004) Brain incorporation of [11C]arachidonic acid, blood volume, and blood flow in health aging: a study with partial volume correction. J. Nucl. Med. 45, 14711479.
  • Greiner R. S., Moriguchi T., Hutton A., Slotnick B. M. and Salem N., Jr (1999) Rats with low levels of brain docosahexaenoic acid show impaired performance in olfactory-based and spatial learning tasks. Lipids 34, S239S243.
  • Hamano H., Nabekura J., Nishikawa M. and Ogawa T. (1996) Docosahexaenoic acid reduces GABA response in substantia nigra neuron of rat. J. Neurophysiol. 75, 12641270.
  • Hamilton L., Greiner R., Salem N., Jr and Kim H. Y. (2000) n-3 Fatty acid deficiency decreases phosphatidylserine accumulation selectively in neuronal tissues. Lipids 35, 863869.
  • Harel Z., Riggs S., Vaz R., White L. and Menzies G. (2001) Omega-3 polyunsaturated fatty acids in adolescents: knowledge and consumption. J. Adolesc. Health 28, 1015.
  • Heird W. C., Prager T. C. and Anderson R. E. (1997) Docosahexaenoic acid and the development and function of the infant retina. Curr. Opin. Lipidol. 8, 1216.
  • Hong S., Gronert K., Devchand P. R., Moussignac R. L. and Serhan C. N. (2003) Novel docosatrienes and 17S-resolvins generated from docosahexaenoic acid in murine brain, human blood, and glial cells. Autacoids in anti-inflammation. J. Biol. Chem. 278, 14 67714 687.
  • Horrocks L. (1985) Metabolism and function of fatty acids in brain, in Phospholipids in Nervous Tissues (Eichberg, J., ed.), pp. 173199. John Wiley, New York.
  • Innis S. M. (2000a) Essential fatty acids in infant nutrition: lessons and limitations from animal studies in relation to studies on infant fatty acid requirements. Am. J. Clin. Nutr. 71, 238S244S.
  • Innis S. M. (2000b) The role of dietary n-6 and n-3 fatty acids in the developing brain. Dev. Neurosci. 22, 474480.
  • Innis S. M. and Dyer R. A. (2002) Brain astrocyte synthesis of docosahexaenoic acid from n-3 fatty acids is limited at the elongation of docosapentaenoic acid. J. Lipid Res. 43, 15291536.
  • Jones C. R., Arai T. and Rapoport S. I. (1997) Evidence for the involvement of docosahexaenoic acid in cholinergic stimulated signal transduction at the synapse. Neurochem. Res. 22, 663670.
  • Kang M. J., Fujino T., Sasano H., Minekura H., Yabuki N., Nagura H., Iijima H. and Yamamoto T. T. (1997) A novel arachidonate-preferring acyl-CoA synthetase is present in steroidogenic cells of the rat adrenal, ovary, and testis. Proc. Natl Acad. Sci. USA 94, 28802884.
  • Khair-El-Din T., Sicher S. C., Vazquez M. A., Chung G. W., Stallworth K. A., Kitamura K., Miller R. T. and Lu C. Y. (1996) Transcription of the murine iNOS gene is inhibited by docosahexaenoic acid, a major constituent of fetal and neonatal sera as well as fish oils. J. Exp. Med. 183, 12411246.
  • Kim H. Y., Karanian J. W., Shingu T. and Salem N., Jr (1990) Stereochemical analysis of hydroxylated docosahexaenoates produced by human platelets and rat brain homogenate. Prostaglandins 40, 473490.
  • Lands W. E. M. and Crawford C. G. (1976) Enzymes of membrane phospholipid metabolism, in The Enzymes of Biological Membranes (Martonosi, A., ed.), pp. 385. Plenum, New York.
  • Langelier B., Furet J. P., Perruchot M. H. and Alessandri J. M. (2003) Docosahexaenoic acid membrane content and mRNA expression of acyl-CoA oxidase and of peroxisome proliferator-activated receptor-delta are modulated in Y79 retinoblastoma cells differently by low and high doses of alpha-linolenic acid. J. Neurosci. Res. 74, 134141.
  • Laposata M., Reich E. L. and Majerus P. W. (1985) Arachidonoyl-CoA synthetase. Separation from nonspecific acyl-CoA synthetase and distribution in various cells and tissues. J. Biol. Chem. 260, 11 01611 020.
  • MacDonald J. I. and Sprecher H. (1991) Phospholipid fatty acid remodeling in mammalian cells. Biochim. Biophys. Acta 1084, 105121.
  • Makrides M., Neumann M. A., Byard R. W., Simmer K. and Gibson R. A. (1994) Fatty acid composition of brain, retina, and erythrocytes in breast- and formula-fed infants. Am. J. Clin. Nutr. 60, 189194.
  • Makrides M., Neumann M. A., Jeffrey B., Lien E. L. and Gibson R. A. (2000) A randomized trial of different ratios of linoleic to α-linolenic acid in the diet of term infants: effects on visual function and growth. Am. J. Clin. Nutr. 71, 120129.
  • Marangell L. B., Martinez J. M., Zboyan H. A., Kertz B., Kim H. F. and Puryear L. J. (2003) A double-blind, placebo-controlled study of the omega-3 fatty acid docosahexaenoic acid in the treatment of major depression. Am. J. Psychiat. 160, 996998.
  • Martin R. E. (1998) Docosahexaenoic acid decreases phospholipase A2 activity in the neurites/nerve growth cones of PC12 cells. J. Neurosci. Res. 54, 805813.
  • Masuzawa Y., Sugiura T., Sprecher H. and Waku K. (1989) Selective acyl transfer in the reacylation of brain glycerophospholipids. Comparison of three acylation systems for 1-alk-1′-enylglycero-3-phophoethanolamine, 1-acylglycero-3-phosphoethanolamine and 1-acylglycero-3-phosphocholine in rat brain microsomes. Biochim. Biophys. Acta 1005, 112.
  • Matsumoto K., Morita I., Hibino H. and Murota S. (1993) Inhibitory effect of docosahexaenoic acid-containing phospholipids on 5-lipoxygenase in rat basophilic leukemia cells. Prostaglandins Leukot. Essent. Fatty Acids 49, 861866.
  • McGahon B. M., Murray C. A., Horrobin D. F. and Lynch M. A. (1999) Age-related changes in oxidative mechanisms and LTP are reversed by dietary manipulation. Neurobiol. Aging 20, 643653.
  • Mischoulon D. and Fava M. (2000) Docosahexanoic acid and omega-3 fatty acids in depression. Psychiatr. Clin. North Am. 23, 785794.
  • Moore S. A. (2001) Polyunsaturated fatty acid synthesis and release by brain-derived cells in vitro. J. Mol. Neurosci. 16, 195200; discussion 215–221.
  • Moore S. A., Yoder E., Murphy S., Dutton G. R. and Spector A. A. (1991) Astrocytes, not neurons, produce docosahexaenoic acid (22 : 6 omega-3) and arachidonic acid (20 : 4 omega 6). J. Neurochem. 56, 518524.
  • Moriguchi T., Loewke J., Garrison M., Catalan J. N. and Salem N., Jr (2001) Reversal of docosahexaenoic acid deficiency in the rat brain, retina, liver, and serum. J. Lipid Res. 42, 419427.
  • Murthy M., Hamilton J., Greiner R. S., Moriguchi T., Salem N., Jr and Kim H. Y. (2002) Differential effects of n-3 fatty acid deficiency on phospholipid molecular species composition in the rat hippocampus. J. Lipid Res. 43, 611617.
  • Neuringer M. (2000) Infant vision and retinal function in studies of dietary long-chain polyunsaturated fatty acids: methods, results, and implications. Am. J. Clin. Nutr. 71, 256S267S.
  • Neuringer M., Connor W. E., Lin D. S., Barstad L. and Luck S. (1986) Biochemical and functional effects of prenatal and postnatal omega-3 fatty acid deficiency on retina and brain in rhesus monkeys. Proc. Natl Acad. Sci. USA 83, 40214025.
  • Noble E. P., Wurtman R. J. and Axelrod J. (1967) A simple and rapid method for injecting H3-norepinephrine into the lateral ventricle of the rat brain. Life Sci. 6, 281291.
  • O'Connor D. L., Hall R., Adamkin D. et al. (2001) Growth and development in preterm infants fed long-chain polyunsaturated fatty acids: a prospective, randomized controlled trial. Pediatrics 108, 359371.
  • Otto S. J., De Groot R. H. and Hornstra G. (2003) Increased risk of postpartum depressive symptoms is associated with slower normalization after pregnancy of the functional docosahexaenoic acid status. Prostaglandins Leukot. Essent. Fatty Acids 69, 237243.
  • Pawlosky R. J., Hibbeln J. R., Lin Y., Goodson S., Riggs P., Sebring N., Brown G. L. and Salem N., Jr (2003) Effects of beef- and fish-based diets on the kinetics of n-3 fatty acid metabolism in human subjects. Am. J. Clin. Nutr. 77, 565572.
  • Paxinos G. and Watson C. (1987) The Rat Brain in Stereotaxic Coordinates, 3nd edn. Academic Press, New York.
  • Poling J. S., Karanian J. W., Salem N., Jr and Vicini S. (1995) Time- and voltage-dependent block of delayed rectifier potassium channels by docosahexaenoic acid. Mol. Pharmacol. 47, 381390.
  • Porcellati G., Goracci G. and Arienti G. (1983) Lipid turnover, in Handbook of Neurochemistry (Lajtha, A., ed.), Vol. 5, pp. 277294. Plenum, New York.
  • Price P. T., Nelson C. M. and Clarke S. D. (2000) Omega-3 polyunsaturated fatty acid regulation of gene expression. Curr. Opin. Lipidol. 11, 37.
  • Purdon A. D. and Rapoport S. I. (1998) Energy requirements for two aspects of phospholipid metabolism in mammalian brain. Biochem. J. 335, 313318.
  • Rapoport S. I. (2001) In vivo fatty acid incorporation into brain phospholipids in relation to plasma availability, signal transduction and membrane remodeling. J. Mol. Neurosci. 16, 243261.
  • Rapoport S. I. (2003) In vivo approaches to quantifying and imaging brain arachidonic and docosahexaenoic acid metabolism in vivo. J. Pediat. 143, S26S34.
  • Rapoport S. I., Chang M. C. J. and Spector A. A. (2001) Delivery and turnover of plasma-derived essential PUFAs in mammalian brain. J. Lipid Res. 42, 678685.
  • Reeves P. G., Nielsen F. H. and Fahey G. C., Jr (1993) AIN-93 purified diets for laboratory rodents: final report of the American Institute of Nutrition ad hoc writing committee on the reformulation of the AIN-76A rodent diet. J. Nutr. 123, 19391951.
  • Reinboth J. J., Clausen M. and Reme C. E. (1996) Light elicits the release of docosahexaenoic acid from membrane phospholipids in the rat retina in vitro. Exp. Eye Res. 63, 277284.
  • Ringbom T., Huss U., Stenholm A., Flock S., Skattebol L., Perera P. and Bohlin L. (2001) Cox-2 inhibitory effects of naturally occurring and modified fatty acids. J. Nat. Prod. 64, 745749.
  • Robinson P. J., Noronha J., DeGeorge J. J., Freed L. M., Nariai T. and Rapoport S. I. (1992) A quantitative method for measuring regional in vivo fatty-acid incorporation into and turnover within brain phospholipids: Review and critical analysis. Brain Res. Rev. 17, 187214.
  • Rojas C. V., Greiner R. S., Fuenzalida L. C., Martinez J. I., Salem N., Jr and Uauy R. (2002) Long-term n-3 FA deficiency modifies peroxisome proliferator-activated receptor beta mRNA abundance in rat ocular tissues. Lipids 37, 367374.
  • Rouser G., Fleischer S. and Yamamoto A. (1970) Two dimensional then layer chromatographic separation of polar lipids and determination of phospholipids by phosphorous analysis of spots. Lipids 5, 494496.
  • Salem N., Jr and Nieblylski C. D. (1995) The nervous system has an absolute molecular species requirement for proper function. Mol. Membrane Biol. 12, 131134.
  • SanGiovanni J. P., Berkey C. S., Dwyer J. T. and Colditz G. A. (2000) Dietary essential fatty acids, long-chain polyunsaturated fatty acids, and visual resolution acuity in healthy full term infants: a systematic review. Early Hum. Dev. 57, 165188.
  • Serhan C. N., Hong S., Gronert K., Colgan S. P., Devchand P. R., Mirick G. and Moussignac R. L. (2002) Resolvins: a family of bioactive products of omega-3 fatty acid transformation circuits initiated by aspirin treatment that counter proinflammation signals. J. Exp. Med. 196, 10251037.
  • Shetty H. U., Smith Q. R., Washizaki K., Rapoport S. I. and Purdon A. D. (1996) Identification of two molecular species of rat brain phosphatidylcholine that rapidly incorporate and turn over arachidonic acid in vivo. J. Neurochem. 67, 17021710.
  • Shimizu T. and Wolfe L. S. (1990) Arachidonic acid cascade and signal transduction. J. Neurochem. 55, 115.
  • Sprecher H. (2000) Metabolism of highly unsaturated n-3 and n-6 fatty acids. Biochim. Biophys. Acta 1486, 219231.
  • Stinson A. M., Wiegand R. D. and Anderson R. E. (1991) Recycling of docosahexaenoic acid in rat retinas during n-3 fatty acid deficiency. J. Lipid Res. 32, 20092017.
  • Strokin M., Sergeeva M. and Reiser G. (2003) Docosahexaenoic acid and arachidonic acid release in rat brain astrocytes is mediated by two separate isoforms of phospholipase A2 and is differently regulated by cyclic AMP and Ca2+. Br. J. Pharmacol. 139, 10141022.
  • Su H. M., Keswick L. A. and Brenna J. T. (1996) Increasing dietary linoleic acid in young rats increases and then decreases docosahexaenoic acid in retina but not in brain. Lipids 31, 12891298.
  • Su H. M., Bernardo L., Mirmiran M., Ma X. H., Corso T. N., Nathanielsz P. W. and Brenna J. T. (1999) Bioequivalence of dietary alpha-linolenic and docosahexaenoic acids as sources of docosahexaenoate accretion in brain and associated organs of neonatal baboons. Pediatr. Res. 45, 8793.
  • Sun G. Y. (1977) Metabolism of arachidonate and stearate injected simultaneously into the mouse brain. Lipids 12, 661665.
  • Sun G. Y. and Su K. L. (1979) Metabolism of arachidonoyl phosphoglycerides in mouse brain subcellular fractions. J. Neurochem. 32, 10531059.
  • Suzuki H., Morikawa Y. and Takahashi H. (2001) Effect of DHA oil supplementation on intelligence and visual acuity in the elderly. World Rev. Nutr. Diet 88, 6871.
  • Tully A. M., Roche H. M., Doyle R., Fallon C., Bruce I., Lawlor B., Coakley D. and Gibney M. J. (2003) Low serum cholesteryl ester-docosahexaenoic acid levels in Alzheimer's disease: a case-control study. Br. J. Nutr. 89, 483489.
  • Uauy R., Mena P. and Rojas C. (2000) Essential fatty acids in early life: structural and functional role. Proc. Nutr. Soc. 59, 315.
  • Van Aerde J. E. and Clandinin M. T. (1993) Controversy in fatty acid balance. Can. J. Physiol. Pharmacol. 71, 707712.
  • Ward G. R., Huang Y. S., Bobik E., Xing H. C., Mutsaers L., Auestad N., Montalto M. and Wainwright P. (1998) Long-chain polyunsaturated fatty acid levels in formulae influence deposition of docosahexaenoic acid and arachidonic acid in brain and red blood cells of artificially reared neonatal rats. J. Nutr. 128, 24732487.
  • Williard D. E., Harmon S. D., Kaduce T. L., Preuss M., Moore S. A., Robbins M. E. and Spector A. A. (2001a) Docosahexaenoic acid synthesis from n-3 polyunsaturated fatty acids in differentiated rat brain astrocytes. J. Lipid Res. 42, 13681376.
  • Williard D. E., Harmon S. D., Preuss M. A., Kaduce T. L., Moore S. A. and Spector A. A. (2001b) Production and release of docosahexaenoic acid by differentiated rat brain astrocytes. World Rev. Nutr. Diet 88, 168172.
  • Woods J., Ward G. and Salem N., Jr (1996) Is docosahexaenoic acid necessary in infant formula? Evaluation of high linolenate diet in the neonatal rat. Pediatr. Res. 40, 687694.
  • Yavin E., Brand A. and Green P. (2002) Docosahexaenoic acid abundance in the brain: a biodevice to combat oxidative stress. Nutr. Neurosci. 5, 149157.
  • Young R. W. (1976) Visual cells and the concept of renewal. Invest. Ophthalmol. Vis. Sci. 15, 700725.