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Brain Advance Access published online on November 20, 2008
Brain, doi:10.1093/brain/awn295
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© The Author (2008). Published by Oxford University Press on behalf of the Guarantors of Brain. All rights reserved. For Permissions, please email: journals.permissions@oxfordjournals.org

Plasmalogens participate in very-long-chain fatty acid-induced pathology

Pedro Brites, Petra A. W. Mooyer, Leila el Mrabet, Hans R. Waterham and Ronald J. A. Wanders

Laboratory Genetic Metabolic Diseases, Academic Medical Center, University of Amsterdam, Amsterdam, The Netherlands

Correspondence to: Pedro Brites, Academic Medical Center, Lab GMZ F0-224, Meibergdreef 9, 1105AZ Amsterdam, The Netherlands E-mail: p.m.brites{at}amc.uva.nl


   Summary
Top
 Summary
Introduction
 Materials and methods
 Results
 Discussion
 Supplementary material
 Funding
 References
 
Peroxisomes are organelles responsible for multiple metabolic pathways including, the biosynthesis of plasmalogens, a class of phospholipids, and the β-oxidation of very-long-chain fatty acids (VLCFA). Lack of peroxisomes or dysfunction in any of their normal functions is the cellular basis for human peroxisomal disorders. Here we used mouse models to understand and define the biochemical and cellular determinants that mediate the pathophysiological consequences caused by peroxisomal dysfunctions. We investigated the role and effects of cellular plasmalogens and VLCFA accumulation in liver, testis and nervous tissue using Pex7 and Abcd1 knockout (KO) mice. In addition, we also generated a Pex7:Abcd1 double KO mouse to investigate how different peroxisomal dysfunctions modulate cellular function and pathology. We found that plasmalogens function as fundamental structural phospholipids and protect cells from damage caused by VLCFA accumulation. In testis, plasmalogens protect spermatocytes from VLCFA-induced degeneration and apoptosis. In nervous tissue, we found that gliosis, inflammatory demyelination and axonopathy caused by accumulation of VLCFA are modulated by plasmalogens. Our findings demonstrate the importance of normal peroxisomal functioning and allow the understanding of the pathological causality of peroxisomal dysfunctions. Nervous tissue deficient in plasmalogens is more prone to damage, illustrating the importance of plasmalogens in peroxisomal disorders including Zellweger syndrome and X-linked adrenoleukodystrophy.

Key Words: peroxisome; plasmalogen; very-long-chain fatty acids; demyelination; neuropathy

Abbreviations: C26:0, hexacosanoic acid; CMAP, compound muscle action potentials; DKO, double knockout; KO, knockout; MNCV, motor nerve conductance velocity; VLCFA, very-long-chain-fatty acids; WT, wild type; X-ALD, X-linked adrenoleukodystrophy

Received July 10, 2008. Revised October 8, 2008. Accepted October 10, 2008.


   Introduction
Top
 Summary
 Introduction
Materials and methods
 Results
 Discussion
 Supplementary material
 Funding
 References
 
The importance of peroxisomes for the normal functioning of cells, tissues and even organisms is underscored by the causative effects of peroxisomal dysfunction. The recognition of various human disorders with impaired peroxisomal functioning has helped the identification of genes involved in peroxisome formation (Weller et al., 2003Go) and the identification and characterization of the extensive repertoire of functions performed by peroxisomes (Wanders and Waterham, 2006Go). These human peroxisomal disorders vary in the age of onset, clinical presentation, tissues affected and pathology. Knowing that peroxisomes participate in at least eight different metabolic pathways (Wanders and Waterham, 2006Go) it has become a challenging task to unravel the physiological link between a given peroxisomal dysfunction and its pathophysiological consequence.

Biological processes that are affected in peroxisomal disorders include endochondral ossification, neuronal migration and myelination (Faust et al., 2005Go). In order to understand the pathophysiological mechanisms behind the metabolic defects caused by peroxisome deficiencies we, and others, have generated mouse models for several human peroxisomal disorders (Baes and Van Veldhoven, 2003Go; Brites et al., 2004aGo). The Pex7 KO mouse serves as a model for Rhizomelic Chondrodysplasia Punctata (RCDP) (Purdue et al., 1999Go; Brites et al., 2003Go). The genetic inactivation of Pex7 in the Pex7 KO mice impairs the import into peroxisomes of proteins carrying a peroxisomal targeting signal type 2 (PTS2), and leads to defects in plasmalogen biosynthesis, phytanic acid {alpha}-oxidation and VLCFA β-oxidation (Brites et al., 2003Go). Throughout life, Pex7 KO mice have impaired biosynthesis of plasmalogens, a plasmenyl phospholipid characterized and distinctive from other glycerophospholipids by the presence of an {alpha},β-unsaturated ether-bond (also called vinyl ether bond) at the sn-1 position of the glycerol backbone (Brites et al., 2004bGo). Plasmalogens comprise all plasmenyl phospholipids containing ethanolamine or choline in the head group of the glycerol backbone. In mammals, the distribution and composition of plasmalogens varies amongst different tissues: nervous tissues, kidney and testes have relatively high levels of plasmalogens whereas liver has very low amounts of plasmalogens. Nervous tissues, kidney and testes are also characterized by high levels of plasmenyl ethanolamine whereas heart and skeletal muscle have high levels of plasmenyl choline (Brites et al., 2004bGo). In Pex7 KO mice the impaired plasmalogen biosynthesis affects all forms of plasmenyl phospholipids (Brites et al., 2003Go).

Plasmalogens have been implicated in several biological processes where they can affect membrane fluidity, mediate signal transduction and protect against oxidative stress (Wanders and Waterham, 2006Go). Nevertheless, the in vivo role(s) of plasmalogens remains elusive and the usage of mouse models defective in plasmalogens may provide a better understanding of their function. Phenotypically, the Pex7 KO mice display, amongst others, defects in embryonic neuronal migration and endochondral ossification (Brites et al., 2003Go). The impairment in neuronal migration found in Pex7 KO mice is relatively mild when compared with the Pex5 KO mice (Baes et al., 1997Go), a mouse that lacks functional peroxisomes and consequently, all peroxisomal functions. The observation that deficient β-oxidation alone does not affect neuronal migration in mice (Baes et al., 2002Go), suggests that multiple peroxisomal impairments possibly including defects in plasmalogen biosynthesis and β-oxidation, may be required to disrupt and modulate the process of neuronal migration.

Normal peroxisomal metabolism is also required for myelination of the nervous system, given that dysmyelination and demyelination are frequently observed in patients with deficiencies in peroxisomal functions (Faust et al., 2005Go), including X-linked Adrenoleukodystrophy (X-ALD). X-ALD is a complex peroxisomal disorder caused by mutations in the ABCD1 gene and biochemically characterized by the abnormal accumulation of VLCFA (Moser et al., 1984Go; Berger and Gartner, 2006Go). Despite its broad range of clinical presentations and outcomes, X-ALD can be divided into a severe variant presenting with a rapidly developing cerebral inflammatory demyelination, and a milder variant characterized by a slower-developing non-inflammatory axonopathy, also designated Adrenomyeloneuropathy (AMN) (Powers et al., 1992Go, 2000Go; Moser, 1997Go). Several independent mouse models of X-ALD, in which the Abcd1 gene has been disrupted (Abcd1 KO mice), have been generated (Forss-Petter et al., 1997Go; Kobayashi et al., 1997Go; Lu et al., 1997Go), but despite the accumulation of VLCFA in target organs, the mice do not develop the characteristic clinical hallmarks of X-ALD. Although some controversy exists about the function of ALDP, the protein encoded by the ABCD1 gene, and the nature of the β-oxidation defect in X-ALD (McGuinness et al., 2003Go; Moser et al., 2007Go), the hypothesis that VLCFA accumulation, at least in mice, is caused by defective transport of these fatty acids across the peroxisomal membrane is still supported by the fact that ALDP belongs to the ATP-binding cassette (ABC) transporter superfamily (Stefkova et al., 2004Go; Wanders et al., 2007Go) and by the observation of reduced rates of VLCFA β-oxidation in fibroblasts (Kobayashi et al., 1997Go; Lu et al., 1997Go; Pujol et al., 2002Go) and hepatocytes (Kobayashi et al., 1997Go) isolated from Abcd1 KO mice.

To investigate the development of pathophysiological alterations caused by peroxisomal dysfunctions we have generated the Pex7:Abcd1 double KO (DKO) mice. These DKO mice allowed us to investigate the role and the effects of plasmalogens and VLCFA under in vivo conditions during postnatal development. Our findings indicate that plasmalogens modulate the pathology caused by VLCFA accumulation with detrimental consequences for spermatocyte development, myelination and axonal survival.


   Materials and methods
Top
 Summary
 Introduction
 Materials and methods
Results
 Discussion
 Supplementary material
 Funding
 References
 
Animal experiments
Mice were genotyped from genomic DNA isolated from toe clippings using the TissueDirect Multiplex PCR system (GenScript Corporation). Primers for the wild type (WT) Pex7 allele were 5'-TCCCAATCTCGAGAGACAGCCGTGTA-3' and 5'-ATGCACAGTAACACTTGGCCTTTTCATGA-3' amplifying a fragment of 450 bp and primers for the mutant Pex7 allele primers were 5'-CTACGTCTGAACGTCAACGTCGAAAACCCG-3' and 5'-GTAGCGGCTGCACAGCGTGTACCAC-3' amplifying a fragment of 300 bp. Genotyping of the Abcd1 alleles was performed with primers 5'-CACAGCCTCTCTCCTTAAGACC-3', 5'-CTCGTTGTCTAGGCAACTGG-3' and 5'-CTTCTATCGCCTTCTTGACG-3', amplifying a fragment of 217 bp for the WT allele and a fragment of 117 bp for the mutant Abcd1 allele.

Pex7 heterozygous mice (Pex7+/–; genotype Ay+/–) in a Swiss Webster background were crossed with Abcd1 KO mice (Abcd1aa; genotype aa+/+) in a C57BL/6J:129S1 mixed background (The Jackson Laboratory). From the F1 we crossed Abcd1 KO, Pex7 heterozygous males (genotype ay+/–) with double heterozygous females (genotype Aa+/–). Abcd1 KO (genotype ay+/+ or aa +/+) and Pex7:Abcd1 DKO (ay–/– or aa–/–) mice were obtained from the crossing of F2 ay+/– males with F2 aa+/– females. WT (genotype Ay+/+ or AA+/+) and Pex7 KO (genotype Ay–/– or AA–/–) mice were obtained from the crossing of F2 Ay+/– males with F2 Aa+/– females. For the experiments we used F3 mice of both sexes since no significant differences were observed between males and females. The numbers of the mice used for each experiment are given in the different figure legends.

Mice were housed under standard conditions and had free access to standard food (TransBreed diet from Special Diets Services, UK) and acidified water. For tissue harvesting, mice were anesthetized with 100 mg/Kg ketamin and 10 mg/kg xylazine. Isolated organs were snap-frozen in liquid nitrogen and stored at –80°C for further analyses. Experiments and mouse manipulations were approved by the University of Amsterdam Animals Experiments Committee.

Cell culture
Skin-derived fibroblasts were grown from skin biopsies taken from 6 months old mice and cultured in Dulbecco's modified Eagle's medium (DMEM), containing 4.5 g/l glucose and L-glutamine, 20% FBS, 100U penicillin/ml, 100 µg/ml streptomycin, 150 µg/ml fungizone and, supplemented with 25 mM Hepes. All studies were performed between passages 16 and 22.

Biochemical analyses
Tissues and fibroblasts were homogenized in phosphate buffer solution (PBS) by sonication. The homogenates were cleared by centrifugation at 900g for 5 min and protein was measured using the DC Protein Assay kit (Bio-Rad) using BSA as standard. For the measurement of VLCFA and phytanic acid lysate corresponding to 300 µg of protein was used and for the measurement of plasmalogens lysate corresponding to 200 µg of protein was used. All biochemical assays, including β-oxidation in cultured fibroblasts (Wanders et al., 1995Go), measurement of VLCFA and phytanic acid levels using a coupled gas chromatography-mass spectrometry method (Vreken et al., 1998Go) and measurement of plasmalogen levels using gas chromatography (Dacremont and Vincent, 1995Go) were performed as described.

Histological analyses
Pieces of harvested tissues were fixed by immersion in buffered formalin at 4°C for 48 h, processed for paraffin embedding and sectioned on a Leica RM2255 microtome, according to routine procedures. Paraffin sections, 5 µm thick, were deparaffinized in Histoclear II (National Diagnostics), rehydrated using decreasing concentrations of ethanol and used for routine histological or immunohistochemical analyses. Routine histological stainings included H&E, nuclear fast red and luxol fast blue stainings. For immunohistochemical analyses we used the Vectorstain Elite peroxidase ABC kit (Vector Laboratories) according to the manufacture's protocol and 3,3'-diaminobenzidine tetrachloride (DAB; Sigma) as a substrate. Processed sections were counterstained with hematoxylin QS (Vector Laboratories), dehydrated in graded ethanol solutions, cleared in Histoclear II and mounted in DPX (Fluka). For immunohistochemical analyses the following primary antibodies and procedures were used: mouse anti-nestin (1:100; Millipore), rabbit anti-GFAP (1:500; DAKOcytomation) after microwaving in Tris:EDTA solution pH 9.0; rabbit anti-cleaved caspase 3 (1:100; Cell Signaling Technology) after microwaving in citrate solution pH 6.0; rat anti-F4/80 (1:20; Serotec) after microwaving in citrate solution pH 6.0; and, goat anti-MBP (1:100; SantaCruz). All secondary antibodies (Vector Laboratories) were biotinylated, and used at 1:100 dilution. In sciatic nerves, MBP detection was performed as described above but after the secondary antibody, slides were incubated with 1:100 dilution of streptavidin-FITC (DAKOcytomation) and observed under UV light from a deuterium lamp using a 450–490 nm excitation filter and a 510–565 nm emission filter. Slides were examined on a Zeiss Axiophot microscope and photographed using a Leica DFC320 camera.

Western blot analysis
Lysates of cerebrum and cerebellum from 10 months old WT and KO mice (n = 3 per genotype) were prepared by sonication in PBS containing 0.2% Triton X100 and protease inhibitor cocktail (Roche). Protein samples (20 µg) were separated on 12.5% SDS-PAGE gels and transferred onto nitrocellulose membranes. Membranes were blocked with 3% skim dried milk (Fluka) in PBS containing 0.01% Tween20 (w/v) and probed with antibodies against MBP (see above) and mouse anti-2',3'-cyclic nucleotide 3' phosphodiesterase (CNPase, Neomarkers) both at 1:250 dilution in blocking buffer. Mouse anti-β-actin (1:5000, Sigma) was used as a protein loading control. Membranes were developed with BCIP/NBT (Promega) after incubation with alkaline phosphatase-labeled secondary antibodies (Biosource). Blots were scanned and quantification was performed using the AIDA software.

Electrophysiology
Mice were anesthetized with 100 mg/kg ketamin and 10 mg/kg xylazine and placed on a warm pad at a temperature of ~30–34°C. Recordings of compound muscle action potentials (CMAP) were obtained on PowerLab 4/25T (AD instruments) using Chart5 software. Recording needle electrodes were placed subcutaneously in the foot pad and supramaximal stimulation of sciatic nerves was performed distally at the level of the ankle and proximally at the sciatic notch. Conduction velocities were calculated as: (proximal distance—distal distance)/(proximal latency—distal latency), with latencies corresponding to the time lapse between the stimulus and the onset of the CMAP and expressed in meters per second.

Data analysis
Data are expressed as means ± SD of measurements. Statistical comparisons were performed using Wilcoxon Mann-Whitney test, and significance was defined as P < 0.05.


   Results
Top
 Summary
 Introduction
 Materials and methods
 Results
Discussion
 Supplementary material
 Funding
 References
 
Generation and phenotypic characterization of Pex7:Abcd1 double KO
To study the pathophysiological consequences of multiple peroxisomal deficiencies we generated a mutant mouse with a combined defect in both plasmalogen biosynthesis and VLCFA β-oxidation. The Pex7:Abcd1 double KO (hereafter referred to as DKO) was obtained after crossing Pex7 heterozygous males with homozygous Abcd1 KO females. At birth, Pex7 KO and DKO mice were smaller than WT and Abcd1 KO mice, displayed hypotonia and ~50% of Pex7 KO and DKO pups died within 2 days of birth. Weight measurements throughout postnatal development consistently demonstrated the impaired development of Pex7 KO and DKO mice when compared to WT and Abcd1 KO mice (Fig. 1A and B). The body weights of DKO mice at P15 (Fig. 1A) and at 3 months of age (data not shown) were more reduced than those of Pex7 KO mice but after 6 months of age Pex7 and DKO mice had similar body weights (Fig. 1B). A factor contributing to the decreased body weight was the extremely reduced amount of white adipose tissue found upon dissection of Pex7 KO and DKO mice (data not shown). Abnormal movement and posture were characteristics found in DKO mice that were not evident in Pex7 KO mice. Pex7 KO and DKO mice that survived past the weaning age had a life-span between 9 and 14 months of age. During this period, DKO mice started to develop tremors and hindlimb ataxia. These phenotypic differences between Pex7 KO and DKO mice suggested a synergy between the two different biochemical abnormalities and their pathological consequences.


Figure 1
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  		Fig. 1 Development and biochemical presentation of mutant knockout mice. Average body weight measurements in male and female mice (6 ≤ n ≥ 3) of all genotypes at the ages of 15 days old (P15) (A) and 9 months old (B). Pex7 KO mice and DKO mice showed deficits in weight-gain throughout the postnatal development, with the DKO showing a more striking phenotype from P15 to 3 months of age, since they weighed significant less even when compared to Pex7 KO mice. Error bars indicate standard deviation. *P < 0.05 compared to WT mice; #P < 0.05 compared to Abcd1 KO mice; +P < 0.05 compared to Pex7 KO mice. (C) Plasmalogen levels in different tissues were detected as their dimethylacetal (DMA) derivatives of the most abundant aliphatic moieties at sn-1 position, i.e. C16:0 (DMA:C16) and C18:0 (DMA:C18) and expressed as percentages of DMA to the corresponding fatty acid. (D) Phytanic acid levels in different tissues from WT and knockout mice. Very low levels of phytanic acid are found in mice fed a standard chow, and Pex7 KO and DKO show levels similar to that of WT and Abcd1 KO mice. Only upon feeding a chow containing 0.5% phytol (P), do Pex7 KO mice accumulate high amounts of phytanic acid (>500-fold compared to control diet C).

 
Plasmalogen levels in brain, kidney and liver did not differ between Pex7 and DKO mice and were extremely reduced (<5%) when compared to WT or Abcd1 KO mice (Fig. 1C). Levels of phytanic acid were very low and did not differ between the different mice (Fig. 1D) fed the standard diet. These results illustrate that the defect in {alpha}-oxidation has no consequences under standard feeding conditions. Only when mice were are fed a diet supplemented with phytol (the precursor of phytanic acid) was a drastic accumulation of phytanic acid observed (Fig. 1D and (Brites et al., 2003Go).

Synergistic effect on VLCFA accumulation in tissues from DKO mice
To evaluate the effects of the combined deficiency of the Pex7 and Abcd1 mutant alleles we measured β-oxidation in skin-derived fibroblasts from WT and mutant mice (Fig. 2A). Whereas mitochondrial β-oxidation of stearic acid was normal in all mutant cell lines, we found a 40% reduction in the β-oxidation rate of the VLCFA hexacosanoic acid (C26:0) in the cell lines from Abcd1, Pex7 and DKO mice (Fig. 2A). Interestingly, despite having similar rates of hexacosanoic acid β-oxidation, we found that the levels of VLCFA in the skin-derived fibroblasts from DKO mice were significantly increased when compared to fibroblasts from Abcd1 and Pex7 KO mice (Fig. 2B). A synergistic defect in peroxisomal β-oxidation of hexacosanoic acid can be clearly identified when analyzing hexacosanoic acid levels in liver and brain (Fig. 2C). Whereas Pex7 KO mice had normal levels of hexacosanoic acid in testis, liver and brain tissue, DKO mice had increased levels that were even higher than the levels found in Abcd1 KO mice. In spleen and spinal cord, Abcd1 and Pex7 KO mice showed similar hexacosanoic acid accumulation, whereas levels of hexacosanoic acid were higher in DKO mice. Measurement of VLCFA in different DKO mice between 9 and 11 months of age revealed levels similar to the ones found at 6 months of age (data not shown). Taken together, these results show that in DKO mice, the defect in VLCFA β-oxidation at the level of Abcd1 combined with the additional defect in β-oxidation at the level of thiolase greatly impairs the ability of peroxisomes to β-oxidize VLCFA in the different tissues from the DKO mice.


Figure 2
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  		Fig. 2 Impaired peroxisomal β-oxidation and accumulation of VLCFA in fibroblasts and tissues. (A) Mitochondrial β-oxidation of stearic acid (C16:0) and peroxisomal β-oxidation of hexacosanoic acid (C26:0) in skin-derived fibroblasts from WT (n = 6), Abcd1 KO (n = 5), Pex7 KO (n = 5) and DKO (n = 6). All KO mice showed specific impairments in peroxisomal β-oxidation. Error bars indicate standard deviation. *P < 0.05 compared to WT mice. (B) Levels of VLCFA in fibroblasts from the different mice (n ≥ 3 for each genotype). (C) Levels of VLCFA in different tissues. Measurements in testis were performed in P20 mice (n = 3 for all genotypes). Measurements in all other organs were performed in 6-month old WT (n = 5), Abcd1 KO (n = 5), Pex7 KO (n = 4) and DKO (n = 5) mice, showing a tissue-dependent accumulation of VLCFA in for Pex7 KO mice in contrast to the generalized accumulation of VLCFA in DKO mice. The different fatty acids have the same color code as in (B). Error bars indicate standard deviation. *P < 0.05 compared to WT mice; #P < 0.05 compared to Abcd1 KO mice; +P < 0.05 compared to Pex7 KO mice.

 
Pathological alterations in non-nervous tissue
We next sought to evaluate the pathological changes in known target tissues and in tissues where we found a differential accumulation of VLCFA. As shown above, normal levels of VLCFA were detected in livers of Pex7 KO mice, but in livers of DKO mice increased levels of VLCFA were observed which were even higher that in Abcd1 KO mice. Histological analysis of livers from WT and mutant mice did not reveal any major changes but we did find increased fibrosis and infiltration surrounding the portal area of livers from Abcd1 and DKO mice (data not shown).

Since male Pex7 KO mice are infertile (Brites et al., 2004aGo) we evaluated the testicular pathology in the different mouse mutants (Fig. 3). At P21, the first signs of pathology in testes of Pex7 KO mice included a slight disorganization of the seminiferous epithelium with respect to the localization of spermatocytes and round spermatids (Fig. 3C). At the same age the testes of Abcd1 KO mice lacked any obvious signs of pathological alterations (Fig. 3B), but the testes of DKO mice showed a severe pathology with very disorganized seminiferous epithelium lacking round spermatids and an increase in the number of abnormally multinucleated cells within the lumen of the seminiferous tubules (Fig. 3D and E), and these multinucleated cells were present in 53 ± 12% of DKO tubules. We next investigated if the loss of spermatocytes might be mediated by apoptosis. Testes of P21 mice were immunostained with an antibody against cleaved-caspase 3, a marker for apoptosis. Spermatocytes positive for cleaved-caspase 3 were evident in the seminiferous tubules of Pex7 (63 ± 3% of tubules contained cleaved-caspase 3 labeled cells) and DKO mice (76 ± 8% of tubules contained cleaved-caspase 3 labeled cells) (Fig. 3F and G). The multinucleated cells observed in DKO were not stained. The number of spermatocytes positive for cleaved-caspase 3 was increased in DKO mice when compared to Pex7 KO mice (Fig. 3H). At 6 months of age, the effects of the continuous testicular degeneration were evident in Pex7 and DKO mice with seminiferous tubules lacking the entire cellular complement. One characteristic difference in the testes of DKO mice was Leydig cell hyperplasia (Fig. 3L). Even at this age, no apparent changes were observed in testes of Abcd1 KO mice (Fig. 3H).


Figure 3
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  		Fig. 3 Plasmalogens modulate the damaging effects of VLCFA accumulation in testis. (A–D) Histogical analyses of testis from P21 WT (A) Abcd1 KO (B), Pex7 KO (C) and DKO (D) mice stained with nuclear fast red, showing degeneration of seminiferous tubules of DKO mice. Arrowheads in (D) point to multinucleated cells found in the lumen of seminiferous tubules. (E) Quantification of multinucleated cells (≥2 nuclei) per seminiferous tubule of mice from the different genotypes. n = 3 for all genotypes and scale bars are 50 µm. (F–G) Immunohistochemical detection of cleaved-caspase 3 in the seminiferous tubules of P21 Pex7 KO (F) and DKO (G) mice showing an increased number of apoptotic spermatocytes in DKO mice. Arrowheads in (G) point to multinucleated cells. (H) Quantification of cleaved-caspase 3 positive cells in seminiferous tubules of mice from the different genotypes. Slides were counterstained with hematoxylin. Scale bars are 50 µm. (I–L) Histogical analyses of testis from 6-month old WT (I) Abcd1 KO (J), Pex7 KO (K) and DKO (L) mice stained with H&E, showing complete degeneration of the seminiferous tubules of Pex7 KO and DKO mice. Arrowheads in (L) denote the increased numbers of Leydig cells in the testis of DKO mice. Identification of Leydig cells was achieved by staining adjacent sections with an antibody against nestin (data not shown). n = 3 for all genotypes and scale bars are 50 µm.

 
VLCFA-induced astrocytosis and microgliosis in DKO mice
We next studied the impact of VLCFA accumulation and plasmalogen deficiency in the central nervous system of the different mutant mice. Gross histological examination of the CNS in the mutant mice revealed loss and mislocalization of Purkinje cells in the cerebellum of Pex7 and DKO mice (data not shown). Since astrocytosis may serve as a marker for damage or dysfunction within the nervous system, brain and spinal cord sections of 9–11 months old mice were immunostained with an antibody against glial fibrillary acidic protein (GFAP; Fig. 4A–F). Astrocytosis was observed throughout the brain of Pex7 and DKO mice but not in brains of Abcd1 KO mice. In the cerebellum (Fig. 4C and D) and brainstem (Fig. 4E and F), reactive astrocytes could be seen both in grey and white matter. Whereas in the cerebellum the degree of astrocytosis was similar between Pex7 and DKO mice, in the brainstem and spinal cord of DKO mice the number of reactive, ameboid astrocytes was increased when compared to Pex7 KO mice.


Figure 4
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  		Fig. 4 Astrocytosis and microgliosis in the CNS of DKO mice. (A–F) Immunohistochemical detection of GFAP in the cerebellum (A–D) and brainstem (E–F) of 10-month old WT (n = 5; A) Abcd1 KO (n = 3, B), Pex7 KO (n = 4, C and E) and DKO (n = 5, D and F), revealing astrocytosis in cerebellar white matter and granular layer of Pex7 KO and DKO mice. In the brainstem of DKO, astrocytes show a more reactive morphology and are present in increased numbers. Scale bars are 100 µm. (G–J) Immunohistochemical detection of F4/80 in the cerebellum (G and H) and brainstem (I and J) of 10-month old Pex7 KO (n = 4, G and I) and DKO (n = 5, H and J), revealing microgliosis in the cerebellar white matter and brainstem of DKO mice. Scale bars are 100 µm.

 
We next sought to investigate the status of microglia in the brains of the mutant mice. Immunostaining with an antibody against F4/80 revealed the exclusive microglyotic state of brains and spinal cords from DKO mice (Fig. 4G–J). Increased numbers of microglia were found in the cerebellar white matter tracts (Fig. 4H) and throughout the brainstem (Fig. 4J) of DKO. Microgliosis was not observed in brains or spinal cords from Abcd1 KO mice (data not shown) or in the cerebellum and brainstem of Pex7 KO mice (Fig. 4G and I). These findings denote an astrocytic response in nervous tissue deficient in plasmalogens and that VLCFA-induced microgliosis is dependent on plasmalogens.

Effect of plasmalogens in VLCFA-induced demyelination
We next assessed the myelination status of the CNS of mutant mice by immunohistochemistry with an antibody against myelin basic protein (MBP; Fig. 5). At 3 months of age, Abcd1, Pex7 and DKO mice showed normal myelination in the corpus callosum, brainstem, spinal cord (data not shown) and cerebellum (Fig. 5B–D). Strikingly, 9–11 months old DKO showed thinning of the cerebellar white matter tracts with reduced staining for MBP, indicative of demyelination (Fig. 5H). The demyelination in DKO mice, as judged by the intensity of MBP staining in IHC, was detected throughout the CNS but was more apparent in smaller white matter tracts compared to large myelinated areas (e.g. corpus callosum; data not shown). No obvious demyelination or parlor was observed in Abcd1 KO mice (Fig. 5F). In Pex7 KO mice only a slight parlor was evident in the small white matter cerebellar tracts (Fig. 5G). Luxol fast blue stainings confirmed the demyelination in small white matter tracts of DKO mice and showed myelin parlor throughout the corpus callosum and the large cerebellar white matter tracts (Supplementary Fig. 1). Western blot analysis of myelin proteins confirmed the more severe demyelination in DKO mice as judged by the reduced amounts of MBP and CNPase (Fig. 5I and J and Supplementary Fig. 2).


Figure 5
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  		Fig. 5 Age-dependent demyelination caused by VLCFA accumulation. (A–D) Immunohistochemical detection of MBP in the cerebellum of 3-months old WT (n = 3; A) Abcd1 KO (n = 3, B), Pex7 KO (n = 3, C) and DKO (n = 3, D), showing normal myelination in the cerebellum. Slides were counterstained with hematoxylin. Scale bars are 50 µm. (E–F) Immunohistochemical detection of MBP in the cerebellum of 10-months old WT (n = 4; A) Abcd1 KO (n = 4, B), Pex7 KO (n = 3, C) and DKO (n = 5, D), showing demyelination of the cerebellar white matter of the third lobule. Slides were counterstained with hematoxylin. Scale bars are 50 µm. Quantification of MBP (I) and CNPase (J) levels in lysates from cerebrum and cerebellum of 10 months old mice. *P < 0.05.

 
Neuropathy with demyelination and axonal loss in DKO mice
We next assessed the condition of the peripheral nervous system (PNS) in the mutant mice. Immunostaining with an antibody against MBP in sciatic nerves of 10-month-old mice revealed a severe loss of myelinated axons in DKO mice with a thinning of the myelin sheaths in the remaining myelinated fibers (Fig. 6D). Pex7 KO nerves were not as affected as the nerves of DKO mice, although we observed loss of small myelinated axons (Fig. 6C) and decreased thickness of the myelin sheaths (data not shown). In Abcd1 KO mice no major changes were observed (Fig. 6B). The observed changes in the PNS of DKO were age-dependent, since at 3 months of age, sciatic nerves from DKO showed only minor changes that, when compared to aged-matched Pex7 KO nerves (Fig. 6E), consisted of a partial loss of myelinated fibers (Fig. 6F).


Figure 6
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  		Fig. 6 Peripheral neuropathy with demyelination and axonal loss in DKO mice. (A–D) Immunofluorescent detection of MBP in sciatic nerves of 10-months old WT (n = 3; A) Abcd1 KO (n = 3, B), Pex7 KO (n = 3, C) and DKO (n = 3, D), revealing thinner myelin sheaths and axonal loss in DKO mice. (E–F) Immunofluorescent detection of MBP in sciatic nerves of 3 months old Pex7 KO (n = 3, E) and DKO (n = 3, F) showing normal myelination of axons. Scale bar is 10 µm. (G) Representative examples of compound muscle action potentials recordings after stimulation at the sciatic notch of 10-month old WT and mutant mice. Increased latencies were observed in Pex7 KO and DKO mice. (H) Motor nerve conductance velocities (MNCV) in 10-months old mice. Bars represent the average values obtained after bilateral measurements in WT (n = 6), Abcd1 KO (n = 4), Pex7 KO (n = 4) and DKO (n = 5) mice. Error bars indicate standard deviation. *P < 0.05.

 
The consequences of the observed pathological alterations in sciatic nerves of Pex7 and DKO mice were evident when we measured motor nerve conductance velocities (MNCV) in 10 months old mice. The electromyograph traces showed increased latencies of the CMAP in Pex7 and DKO mice (Fig. 6G) with DKO nerves being more affected than Pex7 KO nerves since they also had CMAP with prolonged durations. The calculated MNCV (Fig. 8H) revealed decreases in conductance velocity of 36% in Pex7 KO nerves versus 48% in DKO nerves when compared to WT nerves.


   Discussion
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 Materials and methods
 Results
 Discussion
Supplementary material
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 References
 
In this study our aim was to investigate the consequences of VLCFA accumulation and deficiency of plasmalogens and their interrelationship. We show that the combined inactivation of Pex7 and Abcd1 has functional consequences at both the biochemical and pathophysiological level.

We found that the accumulation of VLCFA in Pex7 KO mice was tissue-dependent with normal VLCFA levels in brain and liver but abnormally high VLCFA levels in kidney (Fig. 2). We hypothesized that in the absence of thiolase from peroxisomes, its function can be taken over by the sterol carrier protein x (SCPx), since SCPx can use both straight and branched chain fatty acids as substrates (Seedorf et al., 1994Go; Wanders et al., 1997Go). This hypothesis is strengthened by the fact that SCPx expression is upregulated during postnatal development (Huyghe et al., 2001Go) and that liver peroxisomes contain high amounts of SCPx whereas kidney peroxisomes have very low amounts of SCPx (Mi et al., 2007Go). Thus, in kidney of Pex7 KO mice the absence of thiolase and SCPx leads to a severe defect in VLCFA β-oxidation but in liver and brain, expression of SCPx is able to compensate for the absence of thiolase. The difference in VLCFA found between brain and spinal cord can also be explained by a differential expression level of SCPx. SCPx expression varies significantly between mouse astrocytes, neurons and oligodendrocytes (Cahoy et al., 2008Go). Different proportions of these cells in brain and spinal cord, coupled with different levels of SCPx could be a contributing factor that results in the variable accumulation of VLCFA in Pex7 KO mice.

The genetic inactivation of Abcd1 in Abcd1 KO mice leads to the accumulation of VLCFA in plasma and tissues and is used as a model for X-ALD (Forss-Petter et al., 1997Go; Kobayashi et al., 1997Go; Lu et al., 1997Go; Pujol et al., 2002Go). The Abcd1 encoded protein, i.e. ALDP, is thought to function as the peroxisomal transporter of VLCFA (Stefkova et al., 2004Go; Wanders et al., 2007Go) and it has been hypothesized that the failure to transport VLCFA into peroxisomes may explain the reduced rate of VLCFA β-oxidation and their subsequent accumulation. Despite the accumulation of VLCFA in nervous tissue Abcd1 KO mice fail to develop the hallmarks of X-ALD and only aged Abcd1 KO mice develop some motor abnormalities reminiscent of AMN (Pujol et al., 2002Go).

In this study, by crossing the mutant Pex7 allele with the mutant Abcd1 allele, we blocked peroxisomal β-oxidation at two different steps, i.e. the transport and the degradation of VLCFA and were able to investigate the consequences and the relationship between VLCFA accumulation and plasmalogen deficiency. The biochemical analyses (Fig. 2) clearly demonstrated that in Pex7:Abcd1 DKO mice there is not only a generalized accumulation of VLCFA in cells and tissues but the extent of VLCFA accumulation is higher than in the Abcd1 KO mice. This supports the conclusion of a synergistic block in peroxisomal β-oxidation in DKO mice rather than a merely additive effect. The compensatory effect of SCPx expression found in Pex7 KO mice (Brites et al., 2003Go; Mi et al., 2007Go) was not observed in DKO since the inactivation of ALDP may impair the transport of VLCFA into peroxisomes and thus block peroxisomal β-oxidation at an earlier step.

We observed marked phenotypic consequences of the VLCFA accumulation in several target organs. Testis from Pex7 and DKO mice showed distinct degenerative alterations. Disorganized seminiferous tubules showing abnormalities in spermatocytes were evident in Pex7 KO mice leading to infertility. This pathology is similar to the one found in plasmalogen-deficient testis of Gnpat KO mice (Rodemer et al., 2003Go). Our findings suggest a crucial role of plasmalogens for normal spermatocyte development and in protecting spermatocytes from damage caused by VLCFA accumulation.

Although it is well established that nervous tissues are enriched in plasmalogens, their function(s) within myelin have remained elusive (Brites et al., 2004bGo). Our results indicate that plasmalogens are not required for nervous tissue myelination per se but they may play a role in myelin remodeling, since aged Pex7 KO mice show thinning of myelin sheaths (Fig. 6; Brites et al., unpublished results). Furthermore, based on the results obtained in our DKO mouse, plasmalogens may also have a protective role against the effects of VLCFA accumulation. In plasmalogen-deficient nervous tissues, the damage caused by VLCFA accumulation was evident and marked by demyelination, axonal loss and reactive gliosis. Recently, Kassmann et al. (2007Go) showed that a mouse model with peroxisome-deficient oligodendrocytes, developed demyelination, axonal loss and inflammation. But, since the lack of peroxisomes in these mice yields abnormalities in all the metabolic functions normally performed by peroxisomes (Wanders and Waterham, 2006Go), it is difficult to establish which metabolic pathways are responsible for the observed pathology. The results described in the present study provide important information on the question of which peroxisomal dysfunctions mediate these neuropathological consequences, highlighting the biosynthesis of plasmalogens and VLCFA β-oxidation as key players for normal nervous system functioning.

From the pathological standpoint, our results on the Pex7:Abcd1 DKO mice and those of the mice with peroxisome-deficient oligodendrocytes (Kassmann et al., 2007Go), recapitulate many aspects of the pathology observed in patients with X-ALD (Powers et al., 1992Go, 2000Go; Powers, 1995Go; Moser, 1997Go), whereas the Abcd1 KO mice do not (Forss-Petter et al., 1997Go; Kobayashi et al., 1997Go; Lu et al., 1997Go; Powers et al., 2005Go). The complexity of the X-ALD disorder emerges from a common defect in VLCFA β-oxidation that can present either as a severe and rapidly progressive inflammatory demyelination or a slow non-inflammatory axonopathy (Powers et al., 1992Go, 2000Go). These different presentations of X-ALD have long been taken to indicate the existence of modifier(s) that may play a role in the pathology (Smith et al., 1999Go). Our results suggest that plasmalogens could be one such modifier, as their deficiency augments the damage to the nervous tissue caused by VLCFA accumulation. Measurement of plasmalogen levels in samples from X-ALD patients is scarce but decreased plasmalogen levels have been described both in red blood cells and in white matter samples of X-ALD patients (Wilson and Sargent, 1993Go; Moser et al., 1999Go; Khan et al., 2008Go). Interestingly, decreases in plasmalogen content have also been found in several neuropathological conditions including Alzheimer's disease, Gaucher's disease, dementia and ischemia (Ginsberg et al., 1998Go; Guan et al., 1999Go; Brites et al., 2004bGo; Goodenowe et al., 2007Go; Moraitou et al., 2008Go) and it has been proposed that plasmalogens may modulate the severity and progression of the disease. Although it is still unknown which mechanism, i.e. impaired biosynthesis or increased degradation of plasmalogens is responsible for the loss of plasmalogens in these neurological disorders or in X-ALD, our results suggest that regardless of the mechanism involved, the deficiency of plasmalogens may exacerbate the pathology.

Taken together, our findings demonstrate the crucial importance of peroxisomes in normal development and highlight the relevance of cellular plasmalogens for normal functioning and their possible involvement in the VLCFA-induced damage characteristic of X-ALD. Plasmalogens have been implicated in a variety of cellular processes suggesting that these phospholipids may have both structural and functional roles in membrane and cellular stability, fluidity and protection (Brites et al., 2004bGo). With the increasing in the number of non-peroxisomal disorders that show abnormalities in plasmalogen content, our study highlights the importance of investigating the role of plasmalogens in modulating pathophysiological changes that may contribute to the progression of the disorder.


   Supplementary material
Top
 Summary
 Introduction
 Materials and methods
 Results
 Discussion
 Supplementary material
Funding
 References
 
Supplementary material is available at Brain online.


   Funding
Top
 Summary
 Introduction
 Materials and methods
 Results
 Discussion
 Supplementary material
 Funding
References
 
European Commission (QLG1-CT-2001-01277); The Association Européenne contre les Leucodystrophies (2006-054C1).


   Acknowledgements
 
We thank M. ten Brink, I. Kop and J. de Vos for technical assistance.


   References
Top
 Summary
 Introduction
 Materials and methods
 Results
 Discussion
 Supplementary material
 Funding
 References
 
Baes M, Gressens P, Baumgart E, Carmeliet P, Casteels M, Fransen M, et al. A mouse model for Zellweger syndrome. Nat Genet (1997) 17:49–57.[CrossRef][Web of Science][Medline]

Baes M, Gressens P, Huyghe S, De NK, Qi C, Jia Y, et al. The neuronal migration defect in mice with Zellweger syndrome (Pex5 knockout) is not caused by the inactivity of peroxisomal beta-oxidation. J Neuropathol Exp Neurol (2002) 61:368–74.[Web of Science][Medline]

Baes M, Van Veldhoven PP. Lessons from knockout mice. I: Phenotypes of mice with peroxisome biogenesis disorders. Adv Exp Med Biol (2003) 544:113–22.[Web of Science][Medline]

Berger J, Gartner J. X-linked adrenoleukodystrophy: clinical, biochemical and pathogenetic aspects. Biochim Biophys Acta (2006) 1763:1721–32.[Medline]

Brites P, Motley AM, Gressens P, Mooyer PA, Ploegaert I, Everts V, et al. Impaired neuronal migration and endochondral ossification in Pex7 knockout mice: a model for rhizomelic chondrodysplasia punctata. Hum Mol Genet (2003) 12:2255–67.[Abstract/Free Full Text]

Brites P, Wanders RJ, Waterham HR. The mouse as a model to understand peroxisomal biogenesis and its disorders. Drug Discovery Today: Disease Models (2004a) 1:193–8.[CrossRef]

Brites P, Waterham HR, Wanders RJ. Functions and biosynthesis of plasmalogens in health and disease. Biochim Biophys Acta (2004b) 1636:219–31.[Medline]

Cahoy JD, Emery B, Kaushal A, Foo LC, Zamanian JL, Christopherson KS, et al. A transcriptome database for astrocytes, neurons, and oligodendrocytes: a new resource for understanding brain development and function. J Neurosci (2008) 28:264–78.[Abstract/Free Full Text]

Dacremont G, Vincent G. Assay of plasmalogens and polyunsaturated fatty acids (PUFA) in erythrocytes and fibroblasts. J Inherit Metab Dis (1995) 18(Suppl 1):84–9.[CrossRef][Web of Science][Medline]

Faust PL, Banka D, Siriratsivawong R, Ng VG, Wikander TM. Peroxisome biogenesis disorders: the role of peroxisomes and metabolic dysfunction in developing brain. J Inherit Metab Dis (2005) 28:369–83.[CrossRef][Web of Science][Medline]

Forss-Petter S, Werner H, Berger J, Lassmann H, Molzer B, Schwab MH, et al. Targeted inactivation of the X-linked adrenoleukodystrophy gene in mice. J Neurosci Res (1997) 50:829–43.[CrossRef][Web of Science][Medline]

Ginsberg L, Xuereb JH, Gershfeld NL. Membrane instability, plasmalogen content, and Alzheimer's disease. J Neurochem (1998) 70:2533–8.[Web of Science][Medline]

Goodenowe DB, Cook LL, Liu J, Lu Y, Jayasinghe DA, Ahiahonu PW, et al. Peripheral ethanolamine plasmalogen deficiency: a logical causative factor in Alzheimer's disease and dementia. J Lipid Res (2007) 48:2485–98.[Abstract/Free Full Text]

Guan Z, Wang Y, Cairns NJ, Lantos PL, Dallner G, Sindelar PJ. Decrease and structural modifications of phosphatidylethanolamine plasmalogen in the brain with Alzheimer disease. J Neuropathol Exp Neurol (1999) 58:740–7.[Web of Science][Medline]

Huyghe S, Casteels M, Janssen A, Meulders L, Mannaerts GP, Declercq PE, et al. Prenatal and postnatal development of peroxisomal lipid-metabolizing pathways in the mouse. Biochem J (2001) 353:673–80.[CrossRef][Web of Science][Medline]

Kassmann CM, Lappe-Siefke C, Baes M, Brugger B, Mildner A, Werner HB, et al. Axonal loss and neuroinflammation caused by peroxisome-deficient oligodendrocytes. Nat Genet (2007) 39:969–76.[CrossRef][Web of Science][Medline]

Khan M, Singh J, Singh I. Plasmalogen deficiency in cerebral adrenoleukodystrophy and its modulation by lovastatin. J Neurochem (2008) 106:1766–79.[Web of Science][Medline]

Kobayashi T, Shinnoh N, Kondo A, Yamada T. Adrenoleukodystrophy protein-deficient mice represent abnormality of very long chain fatty acid metabolism. Biochem Biophys Res Commun (1997) 232:631–6.[CrossRef][Web of Science][Medline]

Lu JF, Lawler AM, Watkins PA, Powers JM, Moser AB, Moser HW, et al. A mouse model for X-linked adrenoleukodystrophy. Proc Natl Acad Sci USA (1997) 94:9366–71.[Abstract/Free Full Text]

McGuinness MC, Lu JF, Zhang HP, Dong GX, Heinzer AK, Watkins PA, et al. Role of ALDP (ABCD1) and mitochondria in X-linked adrenoleukodystrophy. Mol Cell Biol (2003) 23:744–53.[Abstract/Free Full Text]

Mi J, Kirchner E, Cristobal S. Quantitative proteomic comparison of mouse peroxisomes from liver and kidney. Proteomics (2007) 7:1916–28.[CrossRef][Web of Science][Medline]

Moraitou M, Dimitriou E, Zafeiriou D, Reppa C, Marinakis T, Sarafidou J, et al. Plasmalogen levels in Gaucher disease. Blood Cells Mol Dis (2008) 41:196–9.[CrossRef][Web of Science][Medline]

Moser AB, Jones DS, Raymond GV, Moser HW. Plasma and red blood cell fatty acids in peroxisomal disorders. Neurochem Res (1999) 24:187–97.[CrossRef][Web of Science][Medline]

Moser HW. Adrenoleukodystrophy: phenotype, genetics, pathogenesis and therapy. Brain (1997) 120:1485–508.[Abstract/Free Full Text]

Moser HW, Mahmood A, Raymond GV. X-linked adrenoleukodystrophy. Nat Clin Pract Neurol (2007) 3:140–51.[CrossRef][Web of Science][Medline]

Moser HW, Moser AE, Singh I, O’Neill BP. Adrenoleukodystrophy: survey of 303 cases: biochemistry, diagnosis, and therapy. Ann Neurol (1984) 16:628–41.[CrossRef][Web of Science][Medline]

Powers JM. The pathology of peroxisomal disorders with pathogenetic considerations. J Neuropathol Exp Neurol (1995) 54:710–19.[Web of Science][Medline]

Powers JM, DeCiero DP, Ito M, Moser AB, Moser HW. Adrenomyeloneuropathy: a neuropathologic review featuring its noninflammatory myelopathy. J Neuropathol Exp Neurol (2000) 59:89–102.[Web of Science][Medline]

Powers JM, Liu Y, Moser AB, Moser HW. The inflammatory myelinopathy of adreno-leukodystrophy: cells, effector molecules, and pathogenetic implications. J Neuropathol Exp Neurol (1992) 51:630–43.[Web of Science][Medline]

Powers JM, Pei Z, Heinzer AK, Deering R, Moser AB, Moser HW, et al. Adreno-leukodystrophy: oxidative stress of mice and men. J Neuropathol Exp Neurol (2005) 64:1067–79.[Web of Science][Medline]

Pujol A, Hindelang C, Callizot N, Bartsch U, Schachner M, Mandel JL. Late onset neurological phenotype of the X-ALD gene inactivation in mice: a mouse model for adrenomyeloneuropathy. Hum Mol Genet (2002) 11:499–505.[Abstract/Free Full Text]

Purdue PE, Skoneczny M, Yang X, Zhang JW, Lazarow PB. Rhizomelic chondrodysplasia punctata, a peroxisomal biogenesis disorder caused by defects in Pex7p, a peroxisomal protein import receptor: a minireview. Neurochem Res (1999) 24:581–6.[CrossRef][Web of Science][Medline]

Rodemer C, Thai TP, Brugger B, Kaercher T, Werner H, Nave KA, et al. Inactivation of ether lipid biosynthesis causes male infertility, defects in eye development and optic nerve hypoplasia in mice. Hum Mol Genet (2003) 12:1881–95.[Abstract/Free Full Text]

Seedorf U, Brysch P, Engel T, Schrage K, Assmann G. Sterol carrier protein X is peroxisomal 3-oxoacyl coenzyme A thiolase with intrinsic sterol carrier and lipid transfer activity. J Biol Chem (1994) 269:21277–83.[Abstract/Free Full Text]

Smith KD, Kemp S, Braiterman LT, Lu JF, Wei HM, Geraghty M, et al. X-linked adrenoleukodystrophy: genes, mutations, and phenotypes. Neurochem Res (1999) 24:521–35.[CrossRef][Web of Science][Medline]

Stefkova J, Poledne R, Hubacek JA. ATP-binding cassette (ABC) transporters in human metabolism and diseases. Physiol Res (2004) 53:235–43.[Web of Science][Medline]

Vreken P, van Lint AE, Bootsma AH, Overmars H, Wanders RJ, van Gennip AH. Rapid stable isotope dilution analysis of very-long-chain fatty acids, pristanic acid and phytanic acid using gas chromatography-electron impact mass spectrometry. J Chromatogr B Biomed Sci Appl (1998) 713:281–7.[CrossRef][Medline]

Wanders RJ, Denis S, Ruiter JP, Schutgens RB, van Roermund CW, Jacobs BS. Measurement of peroxisomal fatty acid beta-oxidation in cultured human skin fibroblasts. J Inherit Metab Dis (1995) 18(Suppl. 1):113–24.[CrossRef][Web of Science][Medline]

Wanders RJ, Denis S, Wouters F, Wirtz KW, Seedorf U. Sterol carrier protein X (SCPx) is a peroxisomal branched-chain beta-ketothiolase specifically reacting with 3-oxo-pristanoyl-CoA: a new, unique role for SCPx in branched-chain fatty acid metabolism in peroxisomes. Biochem Biophys Res Commun (1997) 236:565–9.[CrossRef][Web of Science][Medline]

Wanders RJ, Visser WF, van Roermund CW, Kemp S, Waterham HR. The peroxisomal ABC transporter family. Pflugers Arch (2007) 453:719–34.[CrossRef][Web of Science][Medline]

Wanders RJ, Waterham HR. Biochemistry of mammalian peroxisomes revisited. Annu Rev Biochem (2006) 75:295–332.[CrossRef][Web of Science][Medline]

Weller S, Gould SJ, Valle D. Peroxisome biogenesis disorders. Annu Rev Genomics Hum Genet (2003) 4:165–211.[CrossRef][Web of Science][Medline]

Wilson R, Sargent JR. Lipid and fatty acid composition of brain tissue from adrenoleukodystrophy patients. J Neurochem (1993) 61:290–7.[Web of Science][Medline]


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