Clustering of mutations in the biotin-binding region of holocarboxylase synthetase in biotin-responsive multiple carboxylase deficiency
Clustering of mutations in the biotin-binding region of holocarboxylase synthetase in biotin-responsive multiple carboxylase deficiencyLucie Dupuis, Alfonso Leon-Del-Rio, Daniel Leclerc, Eric Campeau, Lawrence Sweetman1, Jean-Marie Saudubray2, Gail Herman3, K. Michael Gibson4 and Roy A. Gravel*
McGill University, Departments of Biology, Human Genetics and Pediatrics, Montreal Children's Hospital Research Institute, 4060 Ste-Catherine W., Montreal, Quebec, H3Z 2Z3, Canada, 1University of Southern California School of Medicine, Department of Pediatrics and Pathology, Children's Hospital Los Angeles, 4650 Sunset Blvd., Los Angeles, CA 90027, USA, 2Groupe Hospitalier Necker-Enfants Malades, Department of Pediatrics, 142 de Sevres, Paris, 77030, France, 3Baylor College of Medicine, Department of Molecular and Human Genetics, One Baylor Plaza, Houston, TX 77030, USA and 4University of Texas Southwestern Medical Center and Baylor University Medical Center, Institute of Metabolic Diseases, Department of Neurology, 3812 Elm, Dallas, TX 75226, USA
Received March 7, 1996;Revised and Accepted April 4, 1996
Holocarboxylase synthetase (HCS) catalyses the biotinylation of the four biotin-dependent carboxylases found in humans. A deficiency in HCS results in biotin-responsive multiple carboxylase deficiency (MCD). We have identified six different point mutations in the HCS gene in nine patients with MCD. Two of the mutations are frequent among the MCD patients analyzed. Four of the mutations cluster in the putative biotin-binding domain as deduced from the corresponding Escherichia coli enzyme and consistent with an explanation for biotin-responsiveness based on altered affinity for biotin. The two others may define an additional domain involved in biotin-binding or biotin-mediated stabilization of the protein.
Holocarboxylase synthetase (HCS; EC 6.3.4.10) catalyzes the biotinylation of the four biotin-dependent carboxylases found in humans: the mitochondrial propionyl-CoA carboxylase (PCC), pyruvate carboxylase (PC) and [beta]-methylcrotonyl-CoA carboxylase (MCC) and the cytosolic acetyl-CoA carboxylase (ACC) (1 ). An inherited deficiency of HCS activity results in the neonatal form of multiple carboxylase deficiency (MCD), a rare autosomal recessive disorder. The disease is characterized by a decrease in activity of all four biotin-dependent carboxylases so as to impair gluconeogenesis, fatty acid metabolism and amino acid catabolism (1 ). Affected individuals show metabolic acidosis, organic aciduria and varying degrees of hyperammonemia when they first become symptomatic, usually in the first months of life. Clinical manifestations include skin rashes, seizures and developmental delay followed by coma and death if left untreated (1 ). Both the clinical and biochemical symptoms are dramatically resolved with pharmacological doses of oral biotin (1 ). A milder form of the disease (late-onset MCD) has also been described (2 ). Most cases are caused by a defect in biotinidase, the enzyme responsible for the recycling of biotin, although some appear to be mild cases of HCS deficiency (3 ). This additional form of MCD can also be treated with oral biotin.
HCS catalyzes the ATP-dependent covalent attachment of biotin to the apocarboxylases (1 ). While the four mammalian carboxylases differ substantially in their structure, they share a highly homologous biotin-binding site located near the C-terminus in three of them and near the N-terminus in ACC (4 ). HCS activity is found in both the mitochondria and cytosol (5 ,6 ), which allows for the biotinylation of apocarboxylases in both compartments.
Recently, we (7 ) and Suzuki et al. (8 ) cloned the cDNA encoding human HCS, the former by functional complementation of a birA mutant of E. coli defective in the bacterial `biotin ligase' activity and the latter based on peptide sequence data from the purified bovine HCS. Analysis of the predicted amino acid sequence of human HCS (726 amino acids) showed that it is homologous with bacterial biotin ligases in a segment previously identified as the biotin-binding domain of the E.coli BirA protein by X-ray crystallography (9 ). The BirA protein (321 amino acids) serves as co-repressor of the biotin biosynthetic operon as well as carrying the biotin ligase function. Significantly, mutations in birA mutants, showing an increased requirement for biotin in the growth medium (biotin-auxotrophy) have been localized to the biotin-binding region.
In MCD, mutant HCS has been shown to be associated with a reduced affinity for biotin. Burri et al. (10 ), used partially purified extracts from biotin-starved rats as a source of apocarboxylase for the measurement of HCS activity and determined a KM for biotin by assay of activated carboxylase as a measure of product formation. They obtained increases of up to 350-fold in the KM for biotin in MCD patients. Ghneim et al. (11 ), also reported an increased KM in a neonatal MCD patient by assaying the activation of carboxylases in mitochondrial homogenates of biotin-starved fibroblasts. Recently, Suzuki et al. (8 ), identified two mutations, a single base deletion and a point mutation which may be common in Japanese patients with neonatal MCD. However, neither mutation was located in the biotin-binding region.
In the present report, we have screened more than 95% of the HCS coding sequence of six patients with neonatal MCD to reveal several mutations which cluster in the biotin-binding region.
The HCS cDNA was divided into six overlapping segments (Table 1 ) in order to permit SSCP analysis of relatively small fragments. Two PCR products were obtained for fragment I because the 5' end of HCS mRNA is alternatively spliced (7 ). Fragments V and VI contain the putative biotin-binding region. Sequencing the PCR products that showed band shifts on the SSCP gels resulted in the identification of six mutations, of which four were localized to the biotin-binding region (Table 2 ).
. Summary of mutations found in patients with HCS deficiency
Segment
Primers sequenceb
Sizec (bp)
Restriction enzymed
(positiona)
and fragment sizesc
I (-289-79)
TGAGAATTTACAGAGATCATCCTC/
369 & 223
AluI
TCAGAGTGGAGTCCTGCAAGTGCA
213, 94, 62 and 94, 67, 62
II (-75-465)
GCTTGCAGACCTGGGGATCCTTAT/
541
MboII
ACTCTCCTCCCTTCTCTTTCG
241, 207 &93
III (347-869)
ACCATTGAGTCAGTCAAGTTTGCG/
523
FokI
CTGCAGCCACTGCTCAAGACGCT
254, 140 & 129
IV (724-1218)
GAAGGTGTTGGGCCTGTCTTCATC/
495
NlaIII
ATGTTTCCCAAGCCACTGCATAAG
232, 176 & 87
V (1123-1669)
CTGTGACATGAAACAAGTTCCTGC/
547
HaeIII
GAATGGACCTCACTGCTTCCACGA
171, 165, 125 & 86
VI (1527-2181)
GAGGGAATGTGTGGCTGAGCCCTG/
655
HaeIII
TTACCGCCGTTTGGGGAGGATGAG
261, 178, 118 & 98
VI (1527-2181)
GAGGGAATGTGTGGCTGAGCCCTG/
655
HaeIII
aNumbering as in Leon-Del-Rio et al. (7 ).bSequences of primer pairs given 5' to 3', for sense and antisense respectively.cExpected size of amplified fragments. For segment I, two products sizes are given due to alternative splicing (7 ).dRestriction enzymes for digestion of SSCP samples for analysis with resulting calculated sizes of fragments.
. Summary of mutations found in patients with HCS deficiency
Patient
Reference
Mutations outside the biotin-binding region
Mutations within the biotin-binding region
Unidentified
initials
T647 -> G
T1088 -> A
C1522 -> T
G1553 -> A
G1648 -> A
G1711 -> A
second mutation
(L216R)
(V363D)
(R508W)
(G518E)
(G518E)
(G518E)
VE
X
X
MMC
X
X
PD
X
X
JRi
(23-27)
X
X
MK
XX
HB
a
X
X
MC
(28)
X
X
CP
X
Xb
YL
(29)
XX
aAbstract 220 presented at the Vth Nordic Meeting of Medical Genetics in Laugarvatn, Iceland, August 27-28, 1988.bUncharacterized splice mutation (not analyzed at level of genomic DNA).
. Strategy for PCR-based diagnostics of mutations identified in the human HCS gene
Mutation
Primers
Expected
Enzyme restriction site created by:
Expected size after digestion (bp):
size (bp)
Mutation
Mutation
and primer
Normal
Mutant
T647 -> G
CCGTGGACGGACAACTGTCTCC/
CTGCAGCCACTGCTCAAGACGCT
245
HpaII
245
224 & 21
T1088 -> A
GCTCAAGTCAAGCAATTTTAGCAG/
acCAAGGGTTGTCAGAATCTCTC
61
BbsI
61
36 & 25
C1522 -> T
cactgtggcctgtgttccagCA/
tagaaggagactgaactgtaccta
206
NlaIII
206
182 & 24
aC1522 -> T
CGGCAGACCGAGGGCAAAGCA/
TTACCGCCGTTTGGGGAGGATGAG
681
NlaIII
378, 234
378, 211,
36 & 33
36, 33 & 23
G1553 -> A
GGAATGTGTGGCTGAGCCCTGTCG/
tagaaggagactgaactgtaccta
177
TaqI
177
155 & 22
G1648 -> A
cactgtggcctgtgttccagCA/
tagaaggagactgaactgtaccta
206
NlaIII
206
151 & 55
G1711 -> A
cttaattaatgtgcagatttccct/
tagaaggagactgaactgtaccta
242
SspI
242
181 & 61
aIndicates RT-PCR was used to amplify the region with the mutation.The lower case letters indicate intronic sequence and the upper case letters coding sequence.The double underlined letters indicate the nucleotide was altered to create a new restriction site.The single underlined letters indicate the nucleotide was previously altered for a different diagnostic.
In patients MC, CP and YL, a G1648 -> A mutation (Val550Met) was identified which created an NlaIII restriction site. A 206 bp PCR fragment was amplified from genomic DNA using intronic sense and antisense primers (Table 3 ). When the PCR fragment was digested with NlaIII, a 206 bp fragment was obtained for the normal allele, while fragments of 151 and 55 bp were obtained as a result of the base substitution (Fig. 1 A). MC and CP proved to be heterozygous for the mutation and YL appeared homozygous, although partial or complete gene deletion in one allele cannot be excluded in the latter as the parents were unrelated.
A G1553 -> A mutation (Gly518Glu) was identified in one allele of HB which did not alter a restriction site. Therefore, an oligonucleotide was designed to create a TaqI restriction site in the presence of the mutation (Table 3 ). The altered sense primer contained only exonic sequence while the antisense primer contained intronic sequence. When genomic DNA was amplified by PCR with these primers, a band of 177 bp in length was obtained. Upon digestion of the PCR fragment with TaqI, the normal sequence was not cut whereas the mutation resulted in fragments of 155 and 22 bp (Fig. 1 B). HB was confirmed as heterozygous for the mutation.
A T647 -> G mutation (Leu216Arg) was found in patient VE. The mutation was detected using a modified sense primer to create a single HpaII restriction site in the PCR fragment amplified from genomic DNA (Table 3 ). Two exonic primers were used to amplify a 245 bp fragment. When digested with HpaII, fragments of 224 and 21 bp in length were obtained when the mutation was present. The patient was found to be heterozygous for the mutation (Fig. 1 C).
In patients MMC, PD, JRi and MK, a C1522 -> T transition was identified (Arg508Trp). An oligonucleotide was synthesized that generated an NlaIII restriction site in the presence of the mutation (Table 3 ). PCR products for JRi, MK and PD were generated from genomic DNA templates using an altered sense primer which was partially intronic and an antisense primer containing intronic sequence only. For the normal allele, a fragment of 206 bp in length was obtained when the PCR product was digested with NlaIII compared with fragments of 182 and 24 bp when the mutation was present (Fig. 1 D). PD and JRi were heterozygous for the mutation. MK appeared homozygous for the mutation; however, a partial or complete gene deletion in one allele cannot be excluded due to nonconsanguinity. The mutation was also demonstrated in one allele of MMC using an RT-PCR based diagnostic test as genomic DNA was not available. The two cDNA primers, including an altered sense primer, generated a fragment of 681 bp in length (Table 3 ). When the normal PCR product was cut with the enzyme NlaIII, fragments of 378, 234, 36 and 33 bp were generated (only the two largest fragments could be discerned on the gel). In the presence of the mutation, the 234 bp fragment was cut into fragments of 211 and 23 bp (Fig. 1 E).
A G1711 -> A mutation (Asp571Asn) was identified in PD which created a single SspI restriction site. Two intronic primers were used to generate the 242 bp fragment by PCR using genomic DNA as the template (Table 3 ). The 242 bp fragment was cut into segments of 181 and 61 bp in length in the presence of the mutation (Fig. 1 F).
Our results support the prediction originally made by Sweetman's group, that biotin responsiveness in neonatal MCD arises from mutations affecting the affinity of HCS for biotin (12 ). We observed a cluster of four mutations in the predicted biotin-binding region, of which two are candidates for common mutations. The C1522 -> T (Arg508Trp) mutation accounted for five of 24 of the alleles examined (with one patient homozygous for the mutation), while G1648 -> A (Val550Met) accounted for four of 24 of the alleles (with one patient homozygous for the mutation). Furthermore, we have identified a patient with two mutations outside the biotin-binding region. Although we have not expressed these mutations in an in vitro system, the clustering of the mutations, the occurrence of two of them as common alleles and the absence of all of them in a sampling of normal individuals provide compelling evidence that they are responsible for the disease.
We anticipate that the four mutations in the biotin-binding region of HCS, will account for the high KM for biotin measured in patients with neonatal MCD (10 ,11 ). For example, two of the patient fibroblast lines we studied, JRi and MC, had a reported KM of 346 and 48 nM, respectively, compared with 15 nM for the normal enzyme (10 ). JRi was found to have an Arg508Trp mutation and MC was found to have a Val550Met mutation (in each case, the second mutation has yet to be identified). While it is premature to conclude that these mutations are causative of the elevated KM, their location in the binding biotin region and the conservation of three of the four mutations among human, Paracoccus denitrificans, E.coli, Bacillus subtilis, Salmonella typhimurium, mouse (data not shown) and yeast biotin ligases (Fig. 2 ) is consistent with this notion. A stronger case can be made for Arg508Trp, which has a major impact on both the structure and charge of the side chain, than for Val550Met, which represents a conservative change at a site which is variable in the above species.
Fibroblast lines MMC, VE, MC, EG and HB, originating from HCS-deficient patients, were from the cell bank of the Children's Hospital, Los Angeles, CA. Cell line PD was from the Groupe Hospitalier Necker-Enfants Malades, Paris. Additional cell lines surveyed for the identified mutations were from the Baylor Research Institute, Dallas, TX (YL), the Baylor College of Medicine, Houston, TX (MK) and the cell bank of the Children's Hospital, Los Angeles, CA (JRi and CP). The fibroblast lines were grown in [alpha]-MEM medium containing 20% FCS, added biotin (20 mM) and antibiotics/antimycotics (Gibco-BRL). Control fibroblasts were grown in the same medium (without added biotin) in the presence of antibiotics and 10% FCS.
Taq polymerase was purchased from Perkin-Elmer. The T/A cloning kit used for subcloning of PCR products was from Invitrogen. The Gene Clean Kit was obtained from Bio 101 Inc. and the Wizard Mini-Preps used for plasmid purification were obtained from Promega. Plasmid sequencing was done using the Sequenase kit from United States Biochemicals. The [alpha]-[35S]dATP (12.5 mCi/mmol) was purchased from Dupont. The oligonucleotide primers were synthesized by R. Clarizio of the Montreal Children's Hospital Research Institute Oligonucleotide Synthesis Facility.
Total RNA isolated from fibroblasts was reverse transcribed and the HCS cDNA was amplified in six overlapping segments of 450-650 bp in length by PCR according to the methods described earlier (14 ), except that 0.5 [mu]g of oligonucleotide primers were used and the annealing temperature ranged from 60 to 66oC, depending on the segment being amplified. For PCR prior to SSCP analysis, the concentration of dATP was reduced to 6.25 nM, 12.5 [mu]Ci of [alpha]-[35S]dATP was added and the concentration of dTTP, dGTP and dCTP was changed to 12.5 nM.
SSCP analysis was performed according to Orita et al. (15 ), as modified by Triggs-Raine et al. (14 ), except that the [alpha]-35S-labelled PCR products were digested with various restriction enzymes to generate smaller fragments in order to enhance SSCP sensitivity. The fragments were subjected to electrophoresis in a gel containing 6% acrylamide and 10% glycerol in 1* TBE. The digested and non-digested samples were denatured (100oC, 3 min) prior to loading. As well, an aliquot of each sample was run without prior heating to identify the duplex product. The gels were run at 6 W for 16 h. Fragments that displayed band shifts were sequenced directly. PCR products were sequenced either manually (16 ) or were done at the Sequencing Core Facility of the Canadian Genetic Diseases Network (Ottawa, ON) using an Applied Biosystems automated sequencer.
In order to confirm some of the mutations in genomic DNA, the sequences of the flanking exons were determined by inverse PCR to allow the design of primers for the PCR-based diagnostic tests (17 ). Genomic DNA was digested with different enzymes (AluI, RsaI, TaqI, MseI, MspI or HaeII), ligated with T4 DNA ligase (Gibco) and amplified by PCR using adjacent oligonucleotides priming in opposite direction. The PCR products were purified with Gene Clean and were subcloned in the pCRII vector and transformed into E.coli as per the supplier's protocol (TA Cloning Kit, Invitrogen). The candidate clones were then sequenced.
The identified mutations were confirmed directly in PCR amplification products from genomic DNA or reverse transcribed mRNA. The latter was used if DNA was not available. If the mutation created or destroyed a restriction enzyme site, this site was used to confirm the presence of the mutation. A 15 [mu]l sample of the PCR product was digested with the appropriate restriction enzyme and analyzed by electrophoresis on an 8% acrylamide gel. If the base substitution did not change a restriction site, one was created through the diagnostic strategy. An oligonucleotide primer terminating one nucleotide from the mutation was altered one or two nucleotides from its 3' end so as to generate (or eliminate) a restriction site in combination with the mutation. Cleavage of the PCR product with the corresponding restriction enzyme would reveal the presence or absence of the mutation (14 ). For each mutation, a control digestion was included to ensure the PCR products were completely digested.
We thank Stephen Baird (Sequencing Core Facility of the Canadian Genetic diseases Network, Ottawa) for invaluable sequencing assistance and Beverly Akerman (Montreal Children's Hospital Research Institute) for growing the cell cultures. These studies were supported by a grant from the Medical Research Council of Canada. LD is a recipient of a Scholarship from the Montreal Children's Hospital Research Institute. EC is a recipient of a Scholarship from the Fonds de la Recherche en Santé du Québec. We are grateful to the physicians who provided the cell lines used in this study.
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