Significance Although heme synthesis is ubiquitous, specific regulatory mechanisms couple heme production to cellular demand and environmental conditions. The importance of these regulatory mechanisms is highlighted by clinical variability in porphyrias caused by loss-of-function mutations in heme synthesis enzymes. Heme synthesis is also controlled by the mitochondrial AAA+ unfoldase ClpX, which participates in both heme-dependent degradation of δ-aminolevulinate synthase (ALAS) and ALAS activation. This study reports a human familial mutation in CLPX that contributes to erythropoietic protoporphyria (EPP) by partially inactivating CLPX. Reduced CLPX activity increases ALAS post-translational stability, causing pathological accumulation of protoporphyrin IX (PPIX) in human patients. Our results thus identify an additional gene that promotes PPIX overproduction and EPP and highlight the complex gene network contributing to disorders of heme metabolism.

Abstract Loss-of-function mutations in genes for heme biosynthetic enzymes can give rise to congenital porphyrias, eight forms of which have been described. The genetic penetrance of the porphyrias is clinically variable, underscoring the role of additional causative, contributing, and modifier genes. We previously discovered that the mitochondrial AAA+ unfoldase ClpX promotes heme biosynthesis by activation of δ-aminolevulinate synthase (ALAS), which catalyzes the first step of heme synthesis. CLPX has also been reported to mediate heme-induced turnover of ALAS. Here we report a dominant mutation in the ATPase active site of human CLPX, p.Gly298Asp, that results in pathological accumulation of the heme biosynthesis intermediate protoporphyrin IX (PPIX). Amassing of PPIX in erythroid cells promotes erythropoietic protoporphyria (EPP) in the affected family. The mutation in CLPX inactivates its ATPase activity, resulting in coassembly of mutant and WT protomers to form an enzyme with reduced activity. The presence of low-activity CLPX increases the posttranslational stability of ALAS, causing increased ALAS protein and ALA levels, leading to abnormal accumulation of PPIX. Our results thus identify an additional molecular mechanism underlying the development of EPP and further our understanding of the multiple mechanisms by which CLPX controls heme metabolism.

Porphyrias result from disorders of heme synthesis and are associated with mutations in numerous heme synthetic enzymes (1). The genetic penetrance of porphyria-causing alleles is variable, however (1⇓–3), often due to the presence of diverse environmental factors and the existence of modifier genes. In addition, some patients with porphyria who suffer from cutaneous photosensitivity and abnormal liver function have illnesses of unknown etiology, highlighting our incomplete understanding of this complex condition.

Erythropoietic protoporphyria (EPP) is a disorder characterized by the pathological accumulation of a late heme biosynthetic intermediate, protoporphyrin IX (PPIX), in erythroid cells. Approximately 90% of patients with EPP carry a partial deficiency in FECH (EC 4.99.1.1, OMIM 177000), the gene for the mitochondrial enzyme ferrochelatase, which catalyzes the insertion of iron into PPIX for the final step in heme production. In most cases, EPP is due to cosegregation of a heterogeneous, family-specific deleterious FECH allele in compound heterozygosity with the common, low-expression, c.315–48C FECH allele, which affects the use of a cryptic splice-acceptor site in FECH pre-mRNA (4⇓–6). A second, less common class of EPP results from gain-of-function mutations in ALAS2, the erythroid-specific gene for δ-aminolevulinate synthase (ALAS; EC 2.3.1.37, OMIM 300752), which catalyzes the initial step in heme biosynthesis. These dominant, gain-of-function ALAS2 alleles increase ALA production, which triggers the accumulation of downstream heme precursors, especially PPIX. The proportion of patients with EPP with an ALAS2 mutation is <5% in Europe (4, 5) and <10% in the United States (2). Importantly, genetic analysis fails to detect FECH and ALAS2 mutations in 1–5% of families with a member with EPP (which are mostly homozygous for the WT FECH c.315–48T allele), suggesting involvement of loci other than FECH and ALAS2 in this important metabolic illness (3, 7).

Here, we report the identification of a third mechanism underlying EPP in an affected family. This EPP is promoted by a mutation in CLPX, which encodes an AAA+ (ATPases associated with various cellular activities) protein unfoldase. ClpX is widely conserved among bacteria and in the eukaryotic mitochondrion. ClpX works as a ring-shaped homohexamer and is best understood in its function in a proteasome-like complex with the peptidase ClpP (i.e., the ClpXP ATP-dependent protease). For well-studied examples of ClpXP substrates in bacteria, ClpX recognizes specific short sequence motifs in the substrate protein. ClpX unfolds the tertiary structure of substrate protein by ATP-powered polypeptide translocation through its central pore and then presents this unfolded sequence directly into the ClpP proteolytic chamber (reviewed in ref. 8). Recently, ClpX has been identified as a modulator of heme biosynthesis by activating ALAS (9), and also has been observed to mediate the turnover of ALAS under heme-replete conditions; this turnover requires a heme-binding motif in ALAS (10).

Here, we report the identification and analysis of a dominant heterozygous mutation that disables the ATPase activity of CLPX, causing accumulation of PPIX in red blood cells and promoting the development of EPP symptoms. Our experiments demonstrate how this impaired CLPX activity causes deleterious effects in heme metabolism, organismal physiology, and human health.

Discussion Previous work has shown that ClpX activates ALAS by accelerating incorporation of the essential ALAS cofactor, pyridoxal 5′-phosphate (PLP) (9). Knockdown of one of two ClpX homologs in zebrafish, clpxa, caused an erythropoietic anemia that was chemically complemented by the addition of ALA, indicating that this anemia was due to reduced ALAS activity (9). Furthermore, in this previous study, the CLPXP proteolytic complex was not observed to degrade ALAS2 in vitro. The data reported here and by Kubota et al. (10) suggest that additional factors, such as autoregulatory free heme or collaboration with the Lon protease (19), may be required for CLPX-mediated degradation of ALAS. Based on our present results, we suggest a model that integrates both mechanisms by which ClpX modulates ALAS activity, acceleration of PLP cofactor incorporation (activation), and promotion of ALAS protein turnover (degradation) (Fig. 5C). The balance of these mechanisms in vivo determines whether stimulation or suppression of ALAS activity by CLPX is observed. When CLPX is absent or expressed at only a low level, both ALAS activation and degradation are strongly blocked, causing an accumulation of ALAS with decreased enzymatic activity and thus reduced ALA synthesis, causing anemia (Fig. 5C, Right). In the present study, CLPXWT/GD heterozygous individuals contain CLPXWT/GD heteromeric complexes with reduced ATPase activity and weakened interaction with CLPP. The resulting reduction in ALAS degradation has a stronger phenotypic consequence than possible decreased activation, likely due in part to the weaker interaction of CLPXWT/GD heterohexamers with the CLPP peptidase, resulting in a net increase in ALA and PPIX (Fig. 5C, Middle). Furthermore, it is well established with bacterial ClpXP that the unfolding of some protein substrates requires much more ATP and a higher ClpX-ATPase rate compared with other proteins, and the differences depend on the specific structure of the substrate protein (reviewed in ref. 20). The mixed CLPXWT/GD enzyme may be more defective in the complete ALAS unfolding required for degradation than in the limited unfolding that we proposed is required for activation (9). Thus, when cells contain CLPXWT/GD heterohexamers, active, intact ALAS accumulates. Cellular FECH activity and iron levels are insufficient to convert the resulting excess PPIX to heme, causing cells to accumulate free and zinc-PPIX and leading to symptoms of porphyria. The pedigree of the proband’s family clearly demonstrates the dominant coinheritance of PPIX accumulation with the CLPXGD allele through vertical transmission over two generations (Fig.1 and SI Appendix, Fig. S3). Human dominantly inherited diseases are often characterized by incomplete penetrance and variable presentation of disease symptoms, usually due to differences in the influence of environmental factors and/or modifier genes on different individuals. Our data suggest that the human dominant CLPX disorder behaves in this manner, as observed for the previously reported dominant porphyrias (1). Thus, we have uncovered a new molecular basis for EPP, a defect in the AAA+ unfoldase CLPX that results in aberrant control of the key heme synthetic enzyme ALAS. Given the contribution of ClpX and ClpXP to essential organismal processes such as heme synthesis (9, 10) and mitochondrial respiration (21, 22), it is likely that CLPX mutations contribute to additional mitochondriopathies and metabolic disorders. Causative recessive mutations in CLPP have been identified in Perrault syndrome (23, 24). Because no unfoldase-independent role of ClpP is known or predicted, CLPX activity is likely required for the CLPP function(s) that are perturbed in Perrault syndrome. CLPX mutations also could contribute to sideroblastic anemia by slowing PLP incorporation into ALAS2, phenocopying XLPP caused by ALAS2 mutations that inhibit PLP binding (25⇓⇓–28). Treatment with large doses of vitamin B6 (to yield PLP) might bypass a deficiency in CLPX-stimulated PLP incorporation into ALAS2 in a manner analogous to treatment of patients with XLPP with B6-responsive ALAS2 mutations. Because the genetic variant of EPP reported here shares more similarities with ALAS2-based EPP (XLPP) than with FECH-based EPP, it could be helpful to treat patients with CLPX-based EPP with supplemental iron to convert excess PPIX to heme, as has been beneficial for patients with XLPP with a normal FECH gene (3, 29). The incomplete EPP penetrance and variable clinical symptoms observed among the CLPXGD-carrying individuals may be based in environmental factors. In EPP, whether iron therapy is effective or might even sometimes worsen the disease is a matter of debate (30, 31); however, recent work suggests that iron status can modulate the phenotype both in congenital erythropoietic porphyria (32) and in XLPP (3, 5, 29). Therefore, we speculate that the iron deficiency of the index CLPXGD patient may result from an independent pathological process that contributes to the penetrance of EPP. It is our hope that further understanding of the complex network of enzyme and cofactor interactions controlling heme synthesis will continue to contribute ideas for therapeutic strategies to treat diseases caused by aberrant regulation of heme metabolism.

Materials and Methods Human and Vertebrate Animal Study Approval. The study was approved by local Ethics Committees in accordance with the ethical principles for medical research involving human subjects and its subsequent amendments of the Declaration of Helsinki (R162-16–7 and 145–15-4, French ethical agreement). All family members provided written informed consent, and the European Union Institutional Review Board at INSERM approved the protocols. Vertebrate animal studies were performed in compliance with Institutional Animal Care and Use Committee protocols at Boston Children’s Hospital, Brigham & Women’s Hospital, and MIT. Biochemical Analysis. The diagnosis of EPP was based on biochemical analyses of porphyrins in plasma, feces, and erythrocytes, and on FECH enzymatic assays performed according to the European Porphyria Network (EPNET) guidelines and quality control schemes (porphyria.eu/). Erythroid porphyrins were measured as described previously (33). The percentage of zinc-protoporphyrin was calculated from fluorescence emission spectra of extracted of erythrocyte hemolysates. Porphyrins were extracted as described previously (34) and analyzed by HPLC (35). Intracellular ALA levels were measured by LC-MS/MS (UPLC-Xevo TQMS; Waters). Cells were sonicated in the presence of 1 mM succinylacetone, and samples were pretreated by solid-phase extraction using MCX columns (Waters). PPIX in HEK293T and MEL cells and ALAS activity were quantified as described previously (36, 37). Cell Culture and Metabolite Analysis. Lymphoblastoid cells were obtained from the Biological Resource Center, Hôpital Cochin-APHP (Assistance Publique-Hôpitaux de Paris). They were cultured in RPMI 1640 medium (Gibco) supplemented with 10% FBS, 2% l-glutamine 200 mM, 10 mM Hepes, and antibiotics (100 U/mL penicillin and 100 µg/mL streptomycin). CLPX genotypes were identified by Sanger sequencing (SI Appendix, Fig. S3). DS19 MEL clones were a gift from Arthur Skoultchi (Albert Einstein College of Medicine), and the Biological Resource Center at Hôpital Cochin-APHP generously established the lymphoblastoid cell lines. Mononuclear cells were isolated by Ficoll density-gradient centrifugation (PAA Laboratories) from 60 mL of peripheral whole blood. CD34+ cells were purified using immunomagnetic beads (MACS CD34+ MicroBead Kit; Miltenyi Biotec). CD34+ cells were differentiated into erythroid cells by culture in Iscove’s Modified Dulbecco’s Medium (Gibco) supplemented with 15% BIT 9500 (Stemcell Technologies), 2 U/mL EPO, 100 ng/mL SCF, 10 ng/mL IL-6, and 10 ng/mL IL-3 (Miltenyi Biotec), MEL (DS19 subclone) cells were differentiated by incubation in complete DMEM supplemented with 2% DMSO for 3 d. To assess hemoglobinization, o-dianisidine staining was performed as described previously (36). Immunoprecipitation and Western Blot Analysis. Zebrafish embryos at 24 h postfertilization with eGFP expression were lysed in groups of ∼50 embryos in RIPA buffer/protease inhibitor mixture (Roche) by dounce. The clarified soluble lysate after centrifugation was immunoprecipitated with anti-FLAG agarose beads (Sigma-Aldrich) for 12 h at 4 °C, after which immune complex beads were serially washed with PBST/protease inhibitor mixture (Roche) and eluted in boiling Laemmli loading solution. The eluted immune complex was resolved on SDS/PAGE and visualized by Western blot analysis. The following antibodies were used for Western blot analysis: peroxidase-conjugated anti-FLAG M2 (Sigma-Aldrich), anti-ALAS1 (ab84962; Abcam), anti-ALAS2 (ab184964; Abcam or rabbit anti-human ALAS2; Agios Pharmaceutical), anti-CLPX (AP1067b; Abgent), anti-HSP60 (Santa Cruz Biotechnology), anti-ANT (Santa Cruz Biotechnology), and anti–β-actin (Sigma-Aldrich). CHX Block. Inhibition of translation was carried out with 100 μg/mL CHX (Sigma-Aldrich) as described previously (11). Cells were harvested at indicated times after CHX treatment. Proteins obtained from whole-cell lysates (HEK293T cells) or mitochondrial fractions (MEL cells) were resolved by SDS/PAGE. ALAS1, ALAS2, and CLPX were detected by Western blot analysis, and protein levels were quantified using ImageJ (37). CLPX Biochemistry. Murine CLPX variants and human CLPP His6 were expressed and purified as described previously (9). Bacterial expression vectors for the CLPXGD and CLPX-3xFLAG proteins were generated by site-directed mutagenesis and isothermal assembly, respectively, of the previously reported CLPX expression vector. CLPX ATPase activity was monitored at 30 °C by the decrease in absorbance at 340 nm in an NADH-coupled assay (2.5 mM ATP, 1 mM NADH, 7.5 mM phosphoenolpyruvate, 20 U/mL lactate dehydrogenase, and 20 U/mL pyruvate kinase) in 25 mM Hepes pH 7.6, 100 mM KCl, 5 mM MgCl 2 , and 10% glycerol. Co-IP of purified CLPX-3xFLAG and CLPX variants was performed with FLAG M2 antibody-conjugated magnetic agarose beads (Sigma-Aldrich). Here 0.25 μM CLPX-3xFLAG and 0.25 μM CLPX or CLPXGD were incubated in 25 mM Hepes pH 7.6, 100 mM KCl, 5 mM MgCl 2 , 10% glycerol, 0.1% Triton X-100, 0.5 mg/mL acetylated BSA, and 1 mM DTT with 12 μL of beads in a 50-μL total volume on ice for 45 min with intermittent gentle mixing. Beads and bound proteins were washed three times in binding buffer and eluted on ice for 45 min with 30 μL of 1 mg/mL 3xFLAG peptide in binding buffer. Eluted proteins were separated by SDS/PAGE, stained with Sypro Orange, and imaged with a Typhoon FLA 9500 scanner (GE Healthcare). Coisolation of CLPX variants with human CLPP His6 was performed using His-Tag Dynabeads (Thermo Fisher Scientific) in the same buffer used for anti-FLAG co-IP, except for the addition of 10 mM imidazole. CLPX variants were mixed together as indicated, and CLPP His6 and ATPγS were added as indicated. The mixtures were incubated for 10 min at room temperature. His-Tag Dynabeads were added in an equal volume of buffer to yield a concentration of 0.3 μM total CLPX hexamer (CLPX-3xFLAG, CLPXGD, or an equimolar mixture of both) and 0.3 μM CLPP 14-mer in 30 μL. The mixtures were incubated for 10 min at room temperature with intermittent gentle mixing. Beads and bound proteins were washed three times with binding buffer, then eluted for 10 min at room temperature in 30 μL of binding buffer with 300 mM imidazole. Eluted proteins were separated by SDS/PAGE, stained with Sypro Red, and imaged with a Typhoon FLA 9500 scanner (GE Healthcare).

Acknowledgments We thank the individuals and family members for their participation in this study and Jean-Charles Deybach and Said Lyoumi for insightful suggestions during the preparation of the manuscript. We thank Eva Buys and her crew for zebrafish animal husbandry and Thibaud Lefebvre, Steven Ghazal, and Sylvie Simonin for ALA LC/MS/MS, porphyrin measurements, and biostatistics. In vitro HPLC analyses for porphyrins and ALAS activity were performed at the University of Utah Center for Iron and Heme Disorders (under National Institutes of Health Grant U54 DK110858). Ex vivo HPLC and MS analyses for heme precursors and enzyme activity were performed at the French Center for Porphyrias. This work was supported by grants from The Netherlands Society for Biochemistry and Molecular Biology (Nora Baart Foundation), the RadboudUMC Master thesis prize, the Radboud University Master thesis award, and the Dutch Stomach Liver Bowel Foundation (L.N.v.d.V.); the Public Health and Consumer Protection Directorate Public Health Executive Agency of the European Commission, ANR-GIS Maladies Rares Grant ANR07-MRAR-008-01; and the Laboratoire d'Excellence Gr-Ex, reference ANR-11-LABX-0051 (to H.P., L.G., G.N., Z.K., and C.K.). The labex GR-Ex is funded by the program “Investissements d'Avenir” of the French National Research Agency, reference ANR-11-IDEX-0005-02. Additional support was provided by the Howard Hughes Medical Institute (T.A.B.) and National Institutes of Health Grants F32 DK098866 (to Y.Y.Y.), K01 DK106156 (to Y.Y.Y.), F32 DK095726 (to J.R.K.), R01 GM049224 (to T.A.B.), R01 DK020503 (to J.D.P.), U54 DK083909 (to J.D.P.), R01 DK070838 (to B.H.P.), and P01 HL032262 (to B.H.P.). J.R.K. and T.A.B. are employees of the Howard Hughes Medical Institute.