Peroxisome biogenesis disorders

1 Introduction

Peroxisome biogenesis disorders (PBD) are a group of conditions caused by a partial or generalized defect in peroxisome biogenesis. They encompass two phenotypic groups: 1. the Zellweger spectrum disorders (ZSD) including severe, intermediate and milder forms [previously known respectively as Zellweger syndrome (ZS), Neonatal Adrenoleukodystrophy (NALD) and Infantile Refsum Disease (IRD)] and 2. Rhizomelic Chondrodysplasia Punctata type 1 (RCDP1), as well as the variant phenotypes now being described for both groups. PBD represent the paradigm for metabolic malformation syndromes. In a little over 50 years, an intense, concerted multidisciplinary effort has led to elucidating the fundamental biochemical, pathophysiological and molecular bases for these disorders effectively paving the way for therapeutic interventions and trials.

Peroxisomes are membrane bound organelles derived from the endoplasmic reticulum, numbering up to several hundred per mammalian cell, mostly spherical in shape, up to 1μm in diameter. They are indispensable for normal life and highly conserved throughout evolution amongst most eukaryotes. Peroxisomes were first identified by biochemist Christian DeDuve in the 1960 s, as cytoplasmic particles containing hydrogen peroxide-generating oxidases [1]. Later, the neurologist Hans Zellweger reported a group of children with similar craniofacial dysmorphism and malformations in the brain, eye, liver, and kidney [2]. These cases, initially described under the eponym of “cerebro-hepato-renal-syndrome”, were subsequently termed Zellweger syndrome (ZS) [3]. ZS is considered today the prototype for ZSD. The reported absence of morphologically identifiable peroxisomes in hepatocytes and renal tubular cells of affected individuals using the peroxisomal matrix enzyme marker, catalase, [4] demonstrated a connection between ZS and peroxisome dysfunction. However, the etiological connection was not recognized until the identification of multiple peroxisomal enzymes and their deficiencies in affected patients. Similar findings were later demonstrated in the intermediate and milder ZSD phenotypes and RCDP1, which were also being reported at that time [5–7].

2 Peroxisome biochemistry

There are several hundred peroxisomes in all mammalian cells, each containing more than 50 matrix enzymes required for multiple vital metabolic pathways, including the catabolism and synthesis of important cellular lipids [reviewed in [8]]. The best characterized peroxisomal pathways include β-oxidation of very long chain fatty acids (VLCFA), α-oxidation of methyl-branched phytanic acid and the biosynthesis of ether phospholipid (plasmalogen) (see Table 1). The metabolites of these pathways are routinely screened when a peroxisome biogenesis defect is suspected.

Peroxisomal β-oxidation utilizes a variety of different fatty acid substrates in catabolic and synthetic processes: very long (≥ C22) straight chain saturated and unsaturated fatty acids (VLCFA), (2 S) methyl-branched chain pristanic acid and (25 S) di- and tri-hydroxycholestanoic acids (D/THCA). Docosahexanoic acid (DHA), a critical polyunsaturated omega-3 fatty acid, is synthesized in the peroxisome after β-oxidation of its precursor [9–11]. D/THCA are C27 intermediary bile acids that undergo one step of β-oxidation to form the primary C24 bile acids, cholic and chenodeoxycholic acid. Peroxisomal β-oxidation also participates in the regulation of other complex lipids, including pro-inflammatory molecules such as eicosanoids, leukotrienes, and prostaglandins [12–14] [reviewed in [8]]. Furthermore, peroxisomes are the site of β-oxidation of dicarboxylic acids [15]. The initial oxidation step is performed in peroxisomes by acyl- CoA oxidase 1 (ACOX1 for VLCFA, or ACOX2 for pristanic and D/THCA), generating hydrogen peroxide (hence the name, peroxisome) as a byproduct that is detoxified by peroxisomal catalase [16–19] [reviewed in [20]]. D-bifunctional protein (DBP or HSD17B4) catalyzes the second (hydration) and third (dehydration) step of peroxisomal β-oxidation [21–23]. The final step is catalyzed by thiolase (either 3-oxoacyl-CoA thiolase (ACAA1) or SCPX in humans) [reviewed in [8]]. The peroxisomal fatty acid oxidation system is generally not able to degrade fatty acids to completion, so C16 or shorter fatty acids are transported as an acylcarnitine derivative to mitochondria for complete oxidation [11, 24–27].

3-methyl branched fatty acids (predominantly phytanic acid), because of the position of the methyl group, require a preliminary α-oxidation step in order to undergo β-oxidation [28, 29]. This requires the peroxisomal enzymes, phytanyl-CoA hydroxylase (PHYH), phytanyl-CoA lyase, and pristanal dehydrogenase, to generate pristanic acid [30–33] [reviewed in [8]]. Both pristanic acid and D/THCA are generated as (R) and (S) enantiomers, which require conversion to the (S) form by peroxisomal alpha-methyl-CoA racemase (AMACR) before β-oxidation [34]. Phytanic acid is exclusively dietary in origin and obtained from meats and dairy products of ruminant animals and certain fatty fishes [35, 36]. It accumulates in patients with ZSD and RCDP1, as well as in patients with single enzyme defects in PHYH. For a detailed review, see [37].

The committing steps for ether phospholipid synthesis occur in the peroxisome to generate the alkylglycerol backbone and require the peroxisomal enzymes fatty acyl-CoA reductase (FAR1), dihydroxyacetonephosphate acyltransferase (GNPAT), and alkyl-dihydroxy-acetonephosphate synthase (AGPS). Further metabolism in the ER generates the mature ether phospholipids, the majority of which are plasmalogens and contain a fatty alcohol linked by a vinyl ether bond to the sn-1 position of the glycerol backbone and are enriched in DHA and arachidonic acid (AA) at the sn-2 position [reviewed in [38, 39]]. Plasmalogens are a unique class of membrane glycerophospholipids by virtue of their vinyl ether bond, which provides specialized and critical functions to cellular membranes [40–42]. These properties include unique physical membrane attributes [43], signal transduction through released DHA and AA, as well as the remaining lysophospholipid [44], and the ability to provide oxidant protection to other membrane lipids [reviewed in [38, 39]]. Plasmalogens are enriched in heart and skeletal muscles, kidney, brain, and erythrocytes, whereas liver has very low concentration of plasmalogens [45]. Plasmalogen deficiency causes the RCDP phenotype [46–48]. In ZSD, the phenotype is more complex due to multiple enzyme deficiencies. In addition, peroxisomes contain enzymes involved in the catabolism of D-amino acids (D- amino acid oxidase) and L-lysine (pipecolic acid oxidase), polyamine oxidation, detoxification of glyoxylate (alanine-glyoxylate aminotransferase, AGXT), reactive oxygen species, and xenobiotics. [For a full catalog of peroxisomal enzymes see [8]]. Cholesterol levels tend to be low in PBD patients [49–51], although the underlying mechanism remains to be resolved [52]. Recent studies suggest that peroxisome dysfunction leads to chronic ER stress and consequent dysregulation of the sterol response pathway [53].

The catalog of peroxisomal enzymes continues to increase through the use of bioinformatic searches for proteins containing the conserved targeting signals [54–57], as well as proteomics and mass spectrometry techniques [58, 59]. Most peroxisomal enzymes are unique to peroxisomes, but some are shared with other cellular compartments, including mitochondria and cytosol. These multiple subcellular localizations may be due to the presence of multiple targeting signals within the same protein, as shown for 3-hydroxymethylglutaryl-CoA lyase [60], alpha methyl-CoA racemase [61], malonyl-CoA decarboxylase [62] and others, or relate to properties of the oligomeric protein complex as in epoxide hydrolase [63], copper-zinc superoxide dismutase [64], and alanine glyoxylate aminotransferase [65].

Shared enzymes and the movement of substrates and products through various cellular compartments position peroxisomes to be versatile organelles that dynamically adapt their number and metabolic functions in response to cellular needs. This is observed in yeast when grown on fatty acid media, by concomitant transcriptional up-regulation of peroxisomal β-oxidation enzymes and peroxisome proliferation. Rodent peroxisomes also proliferate secondary to increased β-oxidation activity and this is controlled by PPARα [66]. PPARα belongs to a transcription factor superfamily that binds to, and is activated by, a broad range of fatty acids and is a key regulator of peroxisomal and mitochondrial β-oxidation of fatty acids. Nevertheless, PPARα agonists have thus far not been associated with peroxisome proliferation in humans [67]. However, a metabolic control of peroxisome abundance in humans is expected considering the reduced number and enlargement of peroxisomes observed in fibroblast cultures from both ZSD patients and patients with single enzyme defects in β-oxidation pathways (ACOX1 and DBP) [68]. Human peroxisome numbers can also be increased through a PPARα independent mechanism that may rely on PEX11 activation [reviewed in [69]]. Significantly, the PPARγ co-activator (PGC)-1α, a regulator of cellular energy metabolism and mitochondrial biogenesis, was shown also to stimulate peroxisome biogenesis by upregulating PEX11, as well as other PEXproteins [70].

3 Peroxisome biogenesis

Peroxisome biogenesis encompasses the different processes required to assemble and maintain functional peroxisomes including matrix protein import, synthesis of new organelles, and fission of existing organelles. The coordinated activity of 16 PEX proteins, or peroxins, encoded by their corresponding PEX genes, is required for this process in mammals. There is evidence that peroxisome movement and assembly utilizes the cytoskeleton [71][reviewed in [72]]. Current models of eukaryotic peroxisome biogenesis have been recently reviewed [69, 73–75] and briefly are summarized below with emphasis on mammalian systems (see Fig. 1 for diagram of matrix protein import). Peroxisomal enzymes are synthesized on cytosolic ribosomes and contain one of two peroxisomal targeting signals (PTS) required for their import into the peroxisome [reviewed in [76]]. PTS1, the most common, is a C-terminal tripeptide, -Ser-Lys-Leu or consensus variant thereof [77–79]. PTS2 is a N-terminal nonapeptide found in at least 3 mammalian enzymes: 3-oxoacyl-CoA thiolase (ACAA1), PhyH, and AGPS [54, 80–84]. PTS1 proteins are recognized by the cytosolic receptor, PEX5, which also acts as a chaperone for PTS1 proteins [85–87]. The crystal structure of PEX5 demonstrating the PTS1 binding site has been solved [88]. There are two isomers of PEX5: PEX5S and PEX5L, resulting from differential splicing of the primary transcript of the PEX5 gene [89]. The PTS2 sequence forms a structural motif that is recognized by the cytosolic receptor PEX7 [90]. Comparative homology modeling has identified the PEX7-PTS2- binding groove [54]. In mammals, PEX7 needs to interact with PEX5L for targeting to the peroxisomal membrane [89, 91]. PEX5L and PEX7 receptors form cytosolic complexes with their ligands and efficiently ferry them to the peroxisome membrane [91]. The receptor/cargo complex binds at the peroxisome membrane to the PEX13 and PEX14 docking complex [92–96]. The cargo proteins are subsequently translocated into the matrix, possibly through a transient pore, which assembles at the peroxisome membrane to enable matrix protein translocation, before it is disassembled afterwards and its components recycled for further rounds of protein import [97, 98]. Direct evidence for the existence of a transient import pore was shown by electrophysiological studies, which demonstrated that a Pex5p-Pex14p sub-complex of S. cerevisiae harbors pore forming activity [99]. Through this transient pore, which can widen to 9 nm, peroxisomes can import folded, oligomeric and cofactor bound proteins [99–101]. An alternate emerging model postulates that the peroxisomal import machinery displays a preference for monomeric proteins. In this model: (i) newly synthesized peroxisomal proteins are folded by cytosolic chaperones and released as soluble monomers; (ii) PEX5 binding to these monomers (PTS1) blocks their oligomerization; and finally, (iii) these monomeric cargoes are translocated by a single PEX5 molecule into the matrix of the organelle where oligomerization occurs [102].

After translocation, the cargo dissociates from the PTS-receptors and is released into the peroxisomal lumen [103]. The PTS receptors are released from the peroxisomal membrane for another import cycle or directed to the proteasome for their degradation [104–106]. Translocation of the ligands into the peroxisome matrix is also associated with the integral membrane complex: PEX2, PEX10, and PEX12. PEX2/10/12 are ubiquitin ligase (E3)-like proteins containing RING (really interesting new gene) domains and are involved in facilitating the mono- or polyubuiquitination based PTS receptor recycling and degradation [107]. In humans, PEX2 mono-ubiquitination of PEX5 occurs on a conserved cysteine residue in the N-terminal region, and is mediated by redundant ubiquitin-conjugating (E2)-enzymes (UbcH5a, UbcH5b and UbcH5c) [108, 109]. This mono-ubiquitination could provide a handle for PEX5 recycling back to the cytosol [110, 111]. If the monoubiquitination dependent recycling pathway is impaired, the PTS-receptors enter a polyubiquitination-dependent alternative pathway where PEX10/12 poly-ubiquitinate PEX5 on lysine residues, targeting it for degradation by the 26 S proteasome [111, 112]. In wild type cells there is very little receptor turnover by degradation [112, 113]. For a complete review of PEX5 ubiquitination and dislocation, see [114, 115].

The ‘exportomer’ complex of PEX1, PEX6, and PEX26 is involved in the recycling of the PTS1-receptor, PEX5, and presumably PEX7, back to the cytosol for additional rounds of import [116]. PEX1 and PEX6 are members of the AAA protein family (ATPases Associated with various cellular Activities) that contain one or two AAA cassettes responsible for ATP-binding and ATP hydrolysis. PEX1 and PEX6 each contain two ATPase domains (D1 and D2), and are therefore members of the double-ring ATPase family [reviewed in [117]]. These proteins typically form hexameric rings that change conformation during the ATPase cycle, providing the motion and force required for the variety of cellular functions they perform [118–120]. ATP binding is required for the interaction of PEX1 and PEX6 [121], and association with the peroxisome membrane occurs by interaction of the N-terminal region of PEX6 with PEX26 [122]. PEX5 export, but not import, is an ATP driven process, with the energy of ATP hydrolysis providing the conformational change required in the PEX1/6 complex to pull monoubiquitinated PEX5 out of the peroxisome membrane [123]. Recently, cryo-electron microscopy studies have shown that yeast Pex1/Pex6 ATPases form a unique double-ring structure in which the two proteins alternate around the ring [124–126]. These data shed light on the mechanism and function of this ATPase complex and suggest a role in peroxisomal protein import analogous to that of p97 in ER-associated protein degradation, in which the ATPase extracts poly-ubiquitinated proteins from the ER membrane [discussed in [117, 127]].

Peroxisomes can form de novo or by growth and division of pre-existing peroxisomes. PEX3, PEX16, and PEX19 are required for these processes and for correct localization of peroxisome membrane proteins (PMPs) [128–136]. In the de novo pathway, PEX16 is first incorporated into the ER, followed by PEX3 and other PMPs [137]. This leads to the formation of specialized “peroxisome-like” domains, or pre-peroxisomal vesicles, which can detach from the ER to form peroxisomes [137, 138] [reviewed in [139]]. During peroxisome growth, PEX19, a cytosolic protein, recognizes newly synthesized PMPs via their internal targeting signal motifs and binds them [140]. The complex then docks to PEX3 and PEX16, located in the peroxisomal membrane [141, 142]. In this pathway, PEX19-PEX3-PEX16 mediates import for most PMPs, including metabolite transporters and PEX proteins [143][reviewed in [144]]. Peroxisome fission requires the integral membrane protein PEX11β and shares several components of its division machinery with mitochondria. These include dynamin-like proteins (DLPs), Fis1, Mff and ganglioside-induced differentiation-associated protein 1 (GDAP1) [145–148]. PEX11β is required for membrane elongation, and binds to Fis1, which recruits DLPs [reviewed in [69]]. PEX11β is constitutively expressed in all tissues, whereas its isoform PEX11α is inducible [149, 150], and PEX11γ is constitutively expressed in liver [151]. PEX11γ is involved in peroxisome membrane protrusion [152] and may function in the de novo pathway to mediate the import of peroxisomal matrix proteins into the pre-peroxisomal vesicles that are derived from the ER [153]. The relative contribution of the ER-derived versus fission pathway to the total peroxisome population in vivo, and the detailed mechanisms of ER entry and exit of PMPs are controversially discussed. For a detailed review, see [154–156].

Peroxisome turnover, termed pexophagy, is by general mechanisms related to autophagy of cellular contents, as well as mechanisms selective for peroxisomes [reviewed in [157, 158]]. Deficiencies in various PEX proteins increase pexophagy, including PEX3 [159] and PEX14 [160]. Studies in yeast show that deficiencies in exportomer components Pex1, Pex6, or Pex15 (the yeast ortholog of PEX26) increase pexophagy more than other peroxin deficiencies [161]. The constitutive turnover of peroxisomes is estimated to be around 30% per day [162].

4 Clinical disorders

Biallelic pathogenic mutations in any one of 14 PEX genes cause disorders of peroxisome assembly. These are collectively and classically known as peroxisome biogenesis disorders. No human disease has yet been associated with defects in PEX11α and PEX11γ. Defects in PEX1, PEX2, PEX3, PEX5, PEX6, PEX10, PEX11β, PEX12, PEX13, PEX14, PEX16, PEX19 and PEX26 genes cause Zellweger spectrum disorder (ZSD) including Zellweger syndrome (ZS), Neonatal Adrenoleukodystrophy (NALD), and Infantile Refsum Disease (IRD). These historically distinct phenotypes are now considered different presentations within the same clinical and biochemical spectrum with ZS being the most severe presentation, NALD intermediate and IRD milder [4, 164]. The term ZSD is preferred, given the overlap in individual clinical presentations, and the atypical phenotypes currently being identified by new sequencing technologies. In general, disease severity correlates with patient age at onset of symptoms and the predicted consequence of the mutation on peroxin function, capacity for matrix protein import, residual enzyme functions, and peroxisome numbers. At the cellular level, defects causing ZSD lead to decreased numbers of enlarged peroxisomes that show reduced import of matrix enzymes [165]. These structures are termed peroxisome ‘ghosts’ [165, 166]. Defects in PEX7 cause the distinct phenotype of RCDP1 [167]. Since only PTS2 protein import is disrupted in RCDP1, peroxisome number and morphology are normal and there is deficiency of only a subset of matrix enzymes. A phenotypic spectrum of severity is also present in RCDP1 and correlates to residual plasmalogen levels [46–48]. The overall estimated birth incidence for ZSD is 1/50, 000 [168] and 1/100, 000 for RCDP1 [169]. The prevalence of both ZSD and RCDP1 varies among different populations [170–173]. These disorders are autosomal recessive and panethnic, however founder mutations account for the high frequency of the PEX1-c.[2528G>A] allele (p.[Gly843Asp], or p.G843D), the PEX1-c.[2097_2098insT] allele (p.[Ile700Tyrfs42*]) [174, 175], and the PEX7-c.[875T>A] allele (p.[Leu292*]) [172, 177] in individuals of Northern European heritage. Other founder alleles include PEX10-c.[815-815delCT] in the Japanese population [171], PEX6-c.[802_815del] in French Canadians [170], and PEX12- c.[–26G>A; 102A>T] in individuals of Persian Jewish descent [178] (The PEX12-c.[–26G>A] cis mutation is provided by N. Braverman, personal communication).

5 Diagnosis

Clinical suspicion of PBD requires confirmation by determining peroxisome functions (Table 4). The fundamental problem in ZSD is the inability to generate and maintain functional peroxisomes [4, 5].Consequently, the typical biochemical signature of ZSD is concomitant dysfunction of multiple peroxisomal pathways. This is detected in blood, urine and cells by accumulation of VLCFA, phytanic, pristanic and pipecolic acids, D/THCA and deficiency of plasmlaogens, docosahexanoic and primary bile acids (cholic and chenodeoxycholic acids) [diagnosis reviewed in [261]]. In RCDP1, the PEX7 defect impairs only PTS2 protein import, causing reduced plasmalogens and increased phytanic acid levels. Around 10–15% of patients with ZSD phenotypes have single enzyme defects in fatty acid β-oxidation (ACOX1, DBP, SCPX, AMACR), and a similar number of patients with RCDP phenotypes have single enzyme defects in plasmalogen synthesis (GNPAT, AGPS, FAR1). Thus, it is critical to evaluate several peroxisome functions to tease out true PBD from phenocopies due to single enzyme defects. The fact that patients with single enzyme defects in the β-oxidation pathway have phenotypes identical to ZSD [213, 263] implies that β-oxidation defects contribute in large part to the pathology of ZSD. Similarly, defects in plasmalogen synthesis are directly responsible for the RCDP phenotype [46–48, 236]. Metabolite testing is usually followed by enzymatic confirmation of PBD in fibroblast cultures and/or molecular confirmation of the PEX gene defect.

PEX gene sequencing was previously limited due to the challenge of multiple genes involved[174, 264]. However, with the advent of next generation sequencing, molecular diagnosis has become cost efficient, comprehensive, and has enabled additional genotype-phenotype correlations in atypical patients. Over the past decade, studies on more than 800 patients show that defects in PEX1 represent around 60% of all ZSD alleles [174, 265]. In these reports, 12–30% of ZSD alleles are PEX1-p.G843D, and 5–20% are PEX1-p.[Ile700Tyrfs42*]. About 50% of all RCDP alleles are PEX7-p.[Leu292*]. The high frequency of these alleles is due to a founder effect in persons of Northern European extraction [175, 176]. PEX1-p.G843D is a missense protein with residual functions; the presence of at least one allele predicts an intermediate phenotype, and homozygosity generally results in milder phenotypes [227, 266–268]. The large variations in phenotype severity observed amongst PEX1-p.G843D homozygotes (unrelated, as well as intra-familial) suggest the influence of modifier genes [191, 205].

PEX1-p.[Ile700Tyrfs42*] and PEX7-p.[Leu292*], are null alleles and predict a severe phenotype when homozygous [48, 269]. Overall, bi-allelic mutations in PEX1, 6, 26, 10, 12, and 2 contribute to the majority of ZSD alleles [174]. When previously unknown or uncharacterized variants are identified, functional biochemical studies remain a necessary tool to determine causality and pathogenicity [reviewed in [144, 223]].

In addition to diagnoses of affected individuals, early diagnoses of relative asymptomatic infants will be possible with the advent of newborn screening. The combination of liquid chromatography and tandem mass spectrometry (LC–MS/MS) to detect elevated levels of VLCFAs in newborn blood spots was validated as a diagnostic approach for X-linked adrenoleukodystrophy (X-ALD) [270, 271], a single enzyme/protein defect of peroxisome function in which there is only accumulation of VLCFA [reviewed in [272]]. X-ALD screening was recently included in the US Recommended Uniform Screening Panel [273]. The implications of X-ALD newborn screening include the ability to perform clinical surveillance for early detection and treatment of leukodystrophy in affected males, and counseling for carrier females [274]. Newborn screening for X-ALD should also detect the majority of PBD-ZSD cases that feature elevated blood VLCFA levels, thereby permitting early diagnosis and thus more timely institution of supportive management. It will also enable family counseling, determination of accurate incidence figures and continued expansion of the clinical phenotype as variant patients are identified.

6 Management

There are no curative therapies for PBD. Management is multidisciplinary, symptomatic, and based on surveillance of the multiple systems involved. This includes regular developmental, neurological, auditory, visual, nutritional, and orthopedic assessment, and monitoring of liver and adrenal functions [261, 275]. Vitamin K and other fat soluble vitamin supplements are usually prescribed. Care is often palliative for those with severe disease. Genetic counselling and family support organizations provide crucial assistance to parents in their decision making process. The geneticist, neurologist and nutritionist should be involved early in the management of patients with PBD.

Dietary interventions have included restriction of metabolites that accumulate and replacement of those that are deficient. Reports of these approaches in patients are mostly anecdotal and have not been systematically studied. Furthermore, the extensive inter- and intrafamilial variations in the intermediate and milder ZSD patients requires better understanding of the natural history of these disorders and the establishment of reliable clinical endpoints for future interventional trials.

In a small case series in ZSD, recovery of erythrocyte plasmalogens was achieved by administration of the oral plasmalogen precursor batyl alcohol [276], but this did not improve clinical outcome. Oral bile acid supplementation (chenodeoxycholic and ursodeoxycholic acids) in two infants with severe ZSD reduced C27 bile acid intermediates (D/THCA) and improved hepatobiliary function, but did not change the clinical course [278]. In a third infant with severe ZSD, cholic acid supplementation also improved histology with a decrease in bile duct proliferation and inflammation [277]. An infant with AMACR deficiency presenting with cholestasis has done well on long term cholic acid supplementation with complete suppression of C27 bile acids and improvement in liver histology [279]. However, the few other AMACR patients reported were adults with neurological symptoms attributed to pristanic acid accumulation and improved on a phytanic acid restricted diet [213, 280]. Thus the clinical significance of early intervention with bile acid therapy remains unknown.

Patients have also received DHA supplementation, with controversial clinical benefit. In a small case series, improvements in electroretinograms (ERG) and growth were shown [281]. In a double-blind randomized placebo controlled trial that included 48 patients over one year of study, there were no overall improvements in ERG and growth in the treated group compared to the placebo group [207]. Due to the benefit of dietary phytanic acid restriction in adult Refsum disease [282], and the fact that it is not a difficult diet regimen, this is often prescribed for surviving PBD patients but formal studies have not been done to determine if this improves the disease course.

In two siblings with an intermediate-milder ZSD phenotype, the older child received an hepatocyte transplantation [283] and the younger received a liver transplant [284]. Peroxisome biochemical functions improved, but clinical outcome of the younger sibling was relatively better. Another patient received a liver transplant, and biochemical functions improved but it is not yet clear that the clinical outcome has changed [285].

7 Research tools: mouse models

In order to gain further insight into the pathogenesis of ZSD, mouse models with complete deficiency of Pex2, Pex5, Pex13, and Pex11β have been engineered [286]. These mice recapitulate many aspects of severe ZSD, including the metabolic consequences of peroxisome elimination and abnormalities in brain formation, hypotonia, and early demise. Detailed investigations of these models revealed neuronal migration defects commencing early in the cytogenetic epoch and demonstrated by birth dating experiments using 5-BrdU. Other abnormalities include delayed neuron differentiation and extensive neuronal apoptosis (Pex5, Pex11β). In the cerebellum, there was delayed neuronal layering, increased apoptosis in granular neurons and abnormal morphology of Purkinje cells (Pex2) [287]. In Pex11β null mice, there were reduced numbers of importing peroxisomes and only mild deficiencies of β-oxidation and plasmalogen biosynthesis, challenging the idea that PBD clinical features are directly related to the severity of the metabolic abnormalities [288, 289]. Overall, the null models survive gestation, but die soon thereafter, indicating that severe peroxisome deficiency is compatible with embryonic, but not postnatal life in both mice and humans. To gain further information, conditional Pex5 null mice were engineered to evaluate peroxisome dysfunction in key organs. Targeted elimination of peroxisomes in neural precursor cells results in moderate cortical migration defects that correct postnatally and might reflect migration delay [290]. Peroxisome elimination in hepatocytes results in postnatal arrest of neuron migration, particularly in the cerebellum, which still develops in the postnatal period [290]. Thus, peroxisome metabolism in the liver influences the development of the murine brain. It was suggested that elevated C27 bile acids, which are more hydrophobic than the terminal conjugated forms, may enter the brain and impair cerebellar development [290]. Survival of the mice with selective inactivation of Pex5 in neural cells, although neurodevelopmental defects were mild, resulted in a progressive motor and cognitive impairment with death before 6 months of age [291]. Myelinated axons and glial cells were severely affected in the brain and spinal cord, showing decreased compaction of myelin, loss of myelin and axonal damage. Mice with Pex5 inactivation in oligodendrocytes also developed similar axonal damage and demyelination [292]. Thus, peroxisomes may not be required for myelination, but for the preservation of axonal integrity and maintainence of myelin, and this role may be specific to oligodendrocyte peroxisomes [292].

Considering the interrelationship in mitochondrial and peroxisome functions, and the mitochondrial abnormalities noted in tissues from severe ZSD patients [4], mitochondrial abnormalities were investigated in the liver selective Pex5 null mouse model [293]. In this model, morphologically abnormal mitochondria with decreased respiratory chain complex activity were observed in liver. However, oxidative damage was not observed, possibly due to a compensatory increase in glycolysis and mitochondrial proliferation [293]. The decreased gluconeogenesis and glycogen synthesis resulted in the requirement for additional carbohydrates, increased food intake and lower body weights [294]. AMACR deficient mice show a metabolic adaptation that allows them to enhance excretion of C27 bile acids[295].

A PEX1-p.G843D equivalent knock-in mouse model (Pex1-p.G844D) was recently engineered to mimic the common human mutation, and to generate a model for the intermediate and milder phenoytpes that feature progressive peroxisome dysfunction over time. Pex1-p.G844D homozygous mice exhibit many of the typical ZSD manifestations, including growth retardation, progressive retinopathy, bile acid defects resulting in fatty liver with cholestasis, elevated VLCFA, and decreased plasmalogens [296]. Further characterization of the course and mechanisms of disease pathophysiology are underway.

Mice engineered to study RCDP include Pex7 and Gnpat null mice, and Pex7 and Agps hypomorphic models [297–300]. The similarity of phenotypes amongst these models supports plasmalogen deficiency as the primary etiology for RCDP. Thus, they have been used to explore the normal functions of plasmalogens, which are largely unknown and likely to include tissue specific functions [299, 301]. Studies of these mice show requirements for plasmalogens in cerebellar, lens, and skeletal development, as well as spermatogenesis.

The PBD mouse models have also been used to test potential therapeutic interventions. In the Pex5 null models, supplementation with DHA, beginning in utero, normalizes DHA levels in brain but does not improve the neuronal migration disorder [302]. Bile acid supplementation in Pex2 null mice improves cerebellar abnormalities, implicating the accumulation of C27 bile acid intermediates in the cerebellar toxicity observed postnatally in these mice [303]. Supplementation with alkylglycerol, a plasmalogen precursor, can recover plasmalogen levels in PBD patient fibroblast lines [276]. Early alkylglycerol supplementation in the Pex7 null model improved growth and minimized cataract formation [45]. Although plasmalogen levels recovered and tissue pathology improved in somatic tissues, this was not the case in brain, where only around 1% of control plasmalogens were present after 2 months of treatment, and around 2% at 4 months of treatment.

8 Pharmacological therapies under development

9 Conclusion

Enormous progress has been made in understanding mammalian peroxisome biology, biochemistry, and the molecular defects that underlie PBD. The PBD mouse models have already shown enormous utility in determining the disease pathophysiology, and are poised to contribute additional information in the future. Finally, pharmacological interventions for PBD that improve clinical outcomes may be possible. Additional therapeutic interventions can now be explored. The natural history of PBDs needs to be better defined and accurate clinical endpoints established in order to move forward in future systematic clinical trials.