Molecular basis of RhD-positive/D-negative chimerism in two patients

S.S. Eid1

1Princess Iman Centre for Research and Laboratory Sciences, Department of Haematology, King Hussein Medical Centre, Amman, Jordan.

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Volume 10, Nos 1/2, January / March 2004, Pages  228 - 241
 

SUMMARY This study investigated two patients with Rh chimerism: patient A, a healthy individual, and patient B with myelofibrosis. Flow cytometry studies showed two red blood cell populations of Rh phenotypes R1r and rr at percentages of about 25% and 75% respectively. Normal RhD transcript sequences were found following RT-PCR. Genomic DNA (gDNA) showed normal exon, intron, GATA regions and exon/intron boundary sequences except for a single base change in intron 7 (C®A) of exon 7 in patient A. The major change found in both patients was the absence of RHD exon 9 DNA in gDNA isolated from peripheral blood. These findings suggest a somatic mutation, probably in a stem cell common to the myeloid lineage of both patients, and indicate that patient A may undergo malignant transformation in the future.

Introduction

Rh blood group and gene complex

The Rhesus (Rh) blood group system plays a key role in immunohaematology and transfusion medicine. The Rh antigens are the most immunogenic red blood cell protein antigens in humans. Antigens of the Rh blood group system are carried on two proteins encoded by genes denoted RHD and RHCE . Recently, it has been established that the Rh locus on chromosome 1p34.3-p36.1 comprises at least two distinct but highly homologous genes, a D gene and a CcEe gene (Figure 1) [1].

The D and CE polypeptides both consist of 417 amino acids, which differ by 35 amino acids as a result of 44 nucleotide substitutions in the coding sequence [2]. Cherif-Zahar et al. first described the intron-exon organization of the 10-exon RHCE gene. The organization of the closely linked and highly homologous RHD gene appears to be similar [3].

Genetic basis of the RH locus polymorphism

The RH locus is highly polymorphic. The structure of the RH locus was first established by studying blood samples collected from the Caucasian population where the RHD gene is completely deleted in a D-negative phenotype [1]. RHD gene deletion accounts for almost all D-negative pheno- types [1,4]. An intact but dysfunctional RHD gene was reported in a small number of phenotypically D-negative Caucasians. Two examples of such individuals have been studied at the molecular level. Avent et al. [4] reported a nonsense mutation in the RHD gene, while Andrews et al. [5] repor-ted a four-nucleotide deletion in exon 4 of the RHD gene. In the African population a significantly higher proportion (up to 60%) of serologically D-negative individuals carry RHD genes compared with Europeans [6]. Among Japanese people that are typed as D-negative by standard serology, two different RH genotypes can be defined. The first group of individuals lack RHD genes (that is, are genotypically similar to Caucasian D-negatives) and the second group possesses RHD genes. Two groups of workers reported that this second Japanese D-negative allele appears to be of Del (D-elute) phenotype, which can only be identified by complicated adsorption and elution tests [7,8]. However, Okuda et al. [9] stated that this group does not correspond to the Del phenotype, and concluded that the RHD gene is highly detectable among Japanese D-negative individuals. Del has recently been correlated with a 1013 bp deletion, including exon 9 [10], in the RHD gene.

Rh mosaicism and myeloproliferative disorders

Disease-related abnormal expression of blood group antigens has been recognized for a long time. Rh group changes characterized by the presence of two populations of red cells with different phenotypes (Rh mosaicism) have been reported in some patients suffering from acute or chronic myelogenous leukaemia, myeloid metaplasia, polycythaemia and myelofibrosis [11–14]. The myeloproliferative disorders are thought to have a clonal origin arising from a mutation in the haematopoietic pluripotential stem cell [15]. Occasionally the clone has an associated chromosome anomaly or a change in antigenic characteristics. Cooper et al. thought that these changes might also have a clonal origin [16].

Although in some cases there was an association of Rh loss with chromosome aberrations [16–18], no detectable abnormality of chromosome 1, where the RH locus is located (1p34-p36), has been noticed in other cases. In these examples the Rh mosaicism most probably resulted in the expression of an abnormal clone of stem cells (somatic mutation), which occasionally disappeared during clinical remission with a return to a normal Rh phenotype [18,19]. However, it is not clear whether the leukaemic process itself causes these changes in Rh blood group expression or not. Rh mosaicism was also found in apparently healthy individuals in whom chimerism could be eliminated as a possible explanation [13,20,21]. In one case a somatic mutation affecting only one of monozygotic twins was suspected [22]. In a healthy donor and a patient suffering from a non-haematological disease (prolapse of an invertebral disc) a mosaicism for the blood group RH and FY locus (chromosome 1q) was noticed [23,24].

In these studies, serological Rh typing established that persons who had initially typed D+ subsequently had mixed field reactions indicating RhD chimerism. Methods for direct detection of the RHD gene were not available when these studies were reported. The subsequent availability of polymerase chain reaction (PCR) for detecting genes encoding Rh proteins has made it possible to demonstrate the RHD gene even when conventional serological methods do not detect D antigen. Although the molecular basis of RH genes has been largely clarified [25], there is currently no information available regarding the molecular alterations causing Rh blood group changes in malignant diseases, except for one report [14] which studied the molecular basis of RH chimerism in two patients who were about 75% RhD-negative and 25% RhD-positive. One patient suffered from chronic myeloid leukaemia and the other was a normal patient whose Rh chimerism was detected on preoperative blood typing. Both patients were found to have RHCE and not RHD at exon 9.

Methods

Patients

Patient A was a woman aged 25 years old with no haematological disorders or other malignancies. She was found to have Rh chimerism after preoperative (laminectomy) blood group typing. She had not been transfused and does not have a twin.

Patient B was a 79-year-old Caucasian woman, referred by her general practitioner to the haematology outpatients clinic at Norfolk and Norwich hospital for investigation of persistent mild anaemia and leukocytosis. Her blood film and bone marrow aspirate suggested a diagnosis of myelofibrosis.

Blood samples

Blood samples were sent to the International Blood Group Reference Laboratory, Bristol, by the University of Cambridge Divi- sion of Transfusion Medicine, where serological tests and flow cytometry were performed and both patients were diagnosed with Rh chimarism. The International Blood Group Reference Laboratory supplied DNA and cDNA from common RhD-positive and RhD-negative phenotypes.

Genomic DNA extraction and analysis

Genomic DNA (gDNA) was extracted from peripheral blood as described by Avent and Martin [26 ]. PCR reactions were carried out using gDNA templates derived as previously described. Each PCR reaction mix had a final volume of 50 mL consisting of 2.5 mmol/L MgCl2, 10 mmol/L Tris pH 8.3, 1.25 mmol/L dNTPs, 25 mmol/L diluted stocks of primers, 100 ng gDNA and 2.6 U ExpandTM High Fidelity enzyme mix. The PCR reactions were carried out on a Perkin Elmer-Cetus DNA thermal cycler TC1. The PCR conditions and the sets of primers used in the amplification of exons 1–10 are shown in Table 1 and Table 2 respectively. The PCR products were gel-purified using a Qiaex II kit (Qiagen) following the manufacturer’s ins-tructions. Purified DNA was sequenced using dye-labelled terminator cycle sequencing chemistry on an Applied Biosystems 373A DNA sequencer.

PCR amplification of Rh transcripts

Rh transcripts from two overlapping fragments (exon 1–7 and exon 7–10) were isolated, following RT-PCR on total RNA from peripheral blood reticulocytes using Dynabeads Oligo (dT)25. cDNA was prepared as described by Sambrook et al. [27].

Two sets of primers were used to amplify the Rh transcripts. The first set of primers was used to amplify the region from exon 1 to exon 7 and had the following sequences.

• Exon 1 RHD forward (sense) amplimer: 5´-TCCCCATCATAGTCCCTCTG-3´

• Exon 7 RHD reverse (antisense) amplimer: 5´-AAGGTAGGGGCTGGACAG-3´

The second set of primers was used to amplify the region from exon 7 to exon 10 and had the following sequences:

• Exon 7 RHD forward (sense) amplimer: 5´-TGGTGCTTGATACCGCGGAG-3´

• Exon 10 RHD reverse (antisense) amplimer: 5´-AGTGCATAATAAATGGTGAG-3´

PCR reactions were carried out using the following conditions: 94 °C for 1 min, 55 °C for 1 min and 72 °C for 3 min for 35 cycles in a 50 mL reaction mix composed of: 10 mmol/L Tris-HCl pH 8.3, 2.5 mmol/L MgCl2, 1 mm/L each primer, 1.25 mmol/L each dNTP, 100 ng cDNA, and 2.6 U ExpandTM High Fidelity enzyme mix. PCR products were gel purified on 1.5% agarose gel using a Qiaex unit (Qiagen) and cloned into a PCRTM II vector following the manufacturer’s instructions. Sequence analysis of cloned PCR products was performed using dye-labelled terminator cycle sequencing chemistry on an Applied Biosystems 373A DNA sequencer with 0.5 to 1.0 mg plasmid DNA as template. Both strands of DNA were sequenced.

Results

Rh transcript analysis

Reticulocyte RNA isolated from the patient was reverse transcribed and transcripts arising from the RHD gene were amplified using two overlapping sets of primers (exons 1–7 and 7–10). PCR products of the expected sizes (1200 and 387 bp for 1–7, 7–10 respectively) (Figure 2) were cloned into PCRTM II plasmid as described in the Methods. Six clones of each transcript (1–7, 7–10) were isolated and fully sequenced on both strands. The results revealed that all these clones’ sequences are identical to the RHD gene sequence in both patients.

PCR amplification of the gDNA

All gDNA PCR products (Figure 3) were excised, gel-purified, and the two DNA strands sequenced using the same sets of primers used in the amplification. Almost all exons, introns, GATA regions within the promoter region, and exon-intron splicing boundaries were found to be identical to the normal RHD gene. The single exception was intron 7: the primers RHD IN 6F, RHD IN 7R gave no product for patient A, while patient B and the control gave a product with a size of 400 bp (Figures 3 and 4b). When primers RHD EX 7R and RHD 6F were used, there was a product of 3600 bp, which indicates that at least the 5´ half of exon 7D is present. When primers EX 7FOR2 and RH 7R were used to amplify the 3´ regions of exon 7 and intron 7 PCR gave a 3500 bp product (Figure 4a). When sequenced this gave A instead of C (151 exon 7 position) at the 3´ end of primer RHD IN 7R which is located on intron 7 (Figure 5), that is, the CE sequence not the D sequence. This explains why PCR with D IN 6F and D IN 7R failed. When RH CDE IN 7F and RH IN 8R primers were used to amplify exon 8, the PCR gave one product of about 3511 bp for both patients. An interesting finding was that on one occasion, patient A gave two products (Figure 6).

This raises the possibility of a mutation, deletion or insertion in intron 8. When purified, these products gave very poor quality DNA and therefore no sequencing was carried out. The same PCR was repeated more than once. Each time, a single product was obtained (Figure 7a). When EX 9 IN 8F and RH 8/9R primers were used, the PCR gave a 580 bp product (Figure 7b). Patient A and patient B both gave CE-specific sequences only, that is, C at nucleotide 1170 and T at nucleotide 1193 (Figure 8).

Discussion

This study presents the results from two patients, one suffering from a myeloproliferative disorder and the other a healthy individual. Both were females having a mixture of R1r, rr cell populations as demonstrated by serological tests and flow cytometry, which has become a valuable tool in the detection of minority red cell populations.

Loss of RhD antigen in malignant haemopathies and in healthy individuals is extremely rare and molecular information on the blood group changes of these patients is lacking. The molecular basis of Rh chimerism in both patients was studied to find out whether the 2 patients had the same molecular basis, and compare them with those described by Cherif-Zahar et al. (del G600) [14].

The RHD transcripts were studied using two overlapping fragments, exon 1–7 and exon 7–10. The PCR products of both fragments were cloned and sequenced for both patients. As the results indicated, no change was detected in the coding region. However, since the D antigen was not detectable on about 75% of native red cells, either a mutant transcript or truncated protein might be degraded within the cells so that the 7–10 fragment PCR might not amplify immature fragments and pick up only the normal transcript. As a result, no mutant RHD was found because no transcription from the mutant gene occurs. When these PCR amplifications were carried out for all 10 exons of the RHD gene in patients A and B, all exon/intron boundaries, exon, intron, and GATA regions were found to be normal, except for intron 7 in patient A where a single base change was found (C®A) at the 151 exon 7 position, which is at the 3´ end of primer RHD IN 7R, that is, in the CE and not the D sequence. This explains why PCR with RHD IN 6F and RHD IN 7 failed (Figures 3 and 4b). This minor change does not provide an explanation for the patient chimerism, but the presence of CE-specific bases in intron 7 of RHD may suggest that part of the RHD gene in patient A has been replaced by RHCE, resulting in an RHD-CE-D hybrid gene. However, more experiments are needed to see if this is the case.

An alternative explanation is that this change may affect the end part of RHD gene, which results in the loss of RhD antigen. This is supported by the finding that the RHD exon 9 is absent in both patients, with no RHD exon 9 isolated from peripheral blood gDNA. This may be explained by an insertion of DNA (possibly a replacement with part of RHCE) or the deletion of a segment of DNA in intron 8 including exon 9. Any alteration in the amino acid sequence can impair stability, resulting in an unstable molecule that degrades almost as quickly as it is synthesized. As a result no RHD gene is expressed. Possible support for this hypothesis comes from the observation that another primer (D 8/9F, RH IN 8R) produced a large product in high yield (Figure 6a). In patient A, further migration gave two bands in one occasion (Figure 6b), but on other occasions both patients A and B gave only one large product which could not be resolved into discrete bands (Figure 7a). This product needs to be cloned and sequenced to see if any mutation is present in intron 8 which may cause defective processing or splicing of the primary mRNA transcript, resulting in improper translation and the absence of RHD exon 9. When D-specific primers were used (Figure 7a) weak PCR products for RHD exon 9 were produced in patients A and B, stronger in patient B. This may be explained by the high leukocyte count. The presence of these products in both patients when D-specific primers were used may be due to amplification of exon 9 from the minority of cells that are RHD-positive. The high ratio of myeloid cells to reticulocytes may explain why this change could be detected more easily in the gDNA than in the Rh transcripts in patient B, which is not the case for patient A.

Weak D phenotypes are associated with severely depressed D expression. Wagner et al. [28] detected two changes in exon 9: a substitution at nucleotide 1177 (T®G) changing tryptophan to arginine and giving rise to a weak D type 9 phenotype, and another at nucleotide 1154 (G®C) which changes glycine to alanine, and gives rise to a weak D type 2 phenotype. In our patient this is not the case, since the whole RHD exon 9 is absence. In a Japanese population a deletion in 1013 bp in the RHD gene that includes exon 9 has been reported [10]. This deletion is correlated with the Del (D-elute) phenotype (which can only be defined by sophisticated adsorption and elution tests), whereas in our case D antigen expression is severely depressed. Any RHD alteration to exon 9 affects D antigen expression. These findings differ from Cherif-Zahar et al.’s finding [14] of a CML patient whose RhD-positive phenotype shifted to RhD-negative, where sequence analysis of Rh transcripts amplified from reticulocytes revealed a single nucleotide deletion (del G600) localized in a region encoded by exon 4 of the RHD gene.

Comparing the two patients with some of the D-negative phenotypes, it is most likely that the two cases showed a genuine D-negative phenotype caused by clonal changes accompanied by absence of RHD exon 9. More analysis is needed to define the precise mechanism of RhD chimerism, but our results indicate that the defect is within the region of exon 9 in both patients. Northern blotting is helpful in detecting any changes in the RNA level. The 8/9 PCR product (Figure 7a), using primer 1, should be cloned and sequenced. Clinically, the healthy patient has been advised to have regular check-ups to rule out any clonal changes that may develop over time. In the case of the myelofibrosis patient, it may be that during the myelodyplastic process a downregulating gene is activated, inhibiting RHD gene expression.

Acknowledgements

I am most grateful to my supervisor, Dr N. Avent, for his help and guidance throughout this project.

References

1. Colin Y et al. Genetic basis of the RhD-positive and RhD-negative blood group polymorphism as determined by Southern analysis. Blood, 1991, 78:2747–52.

2. Arce MA et al. Molecular cloning of RhD cDNA derived from a gene present in RhD-positive, but not RhD-negative individuals. Blood, 1993, 82:651–5.

3. Cherif-Zahar B et al. Organization of the gene (RHCE) encoding the human blood group RhCcEe antigens and characterization of the promoter region. Genomics, 1994, 19:68–74.

4. Avent ND et al. Evidence of genetic diversity underlying RhD-, weak D (Du), and partial D phenotypes as determined by multiplex polymerase chain reaction analysis of the RHD gene. Blood, 1997, 89:2568–77.

5. Andrews KT et al. The RhD- trait in a white patient with the RhCCee phenotype attributed to a four-nucleotide deletion in the RHD gene. Blood, 1998, 92: 1839–40.

6. Daniels G, Green C, Smart E. Differences between RhD-negative Africans and RhD-negative Europeans. Lancet, 1997, 350:862–3.

7. Fukumori Y et al. Further analysis of Del (D-elute) using polymerase chain reaction (PCR) with RHD gene-specific primers. Transfusion medicine, 1997, 7: 227–31.

8. Sun CF et al. RHD gene polymorphisms among RhD-negative Chinese in Taiwan. Vox sanguinis, 1998, 751:52–7.

9. Okuda H et al. The RHD gene is highly detectable in RhD-negative Japanese donors. Journal of clinical investigation, 1997, 100:373–9.

10. Chang JG et al. Human RhDel is caused by a deletion of 1,013 bp between introns 8 and 9 including exon 9 of RHD gene. Blood, 1998, 92:2602–4.

11. Majsky A. Some cases of leukaemia with modifications of the D(Rho)-receptor. Neoplasma, 1967, 14:335–44.

12. Habibi B, Lopez M, Salmon C. Two new cases of Rh mosaicism. Selective study of red cell populations. Vox sanguinis, 1974, 27:232–42.

13. Mertens G et al. Loss of D and C expression in chronic myelomonocytic leukemia. Transfusion, 1997, 37:880–1.

14. Cherif-Zahar B et al. Shift from Rh-positive to Rh-negative phenotype caused by a somatic mutation within the RHD gene in a patient with chronic myelocytic leukaemia. British journal of haemato-ogy, 1998, 102:1263–70.

15. Barr RD, Fialkow PJ. Clonal origin of chronic myelocytic leukaemia. New England journal of medicine, 1973, 289: 307–9.

16. Cooper B et al. Loss of Rh antigen associated with acquired Rh antibodies and a chromosome translocation in a patient with myeloid metaplasia. Blood, 1979, 54:642–7.

17. Marsh WL et al. Mapping human autosomes: evidence supporting assignment of rhesus to the short arm chromosome no.1. Science, 1974, 183:966–8.

18. Mohandas V et al. Loss and reappearance of Rho(D) antigen in an individual with acute myelogenous leukemia. Transfusion, 1993, 33:24s.

19. Bracey AW et al. Rh mosaicism and aberrant MNSs antigen expression in a patient with chronic myelogenous leukaemia. American journal of clinical pathology, 1983, 79:397–401.

20. Race RR, Sanger R. Blood groups in man. Oxford, Blackwell, 1975:178–260.

21. Salaru NN, Lay WH. Rh blood group mosaicism in a healthy elderly woman. Vox sanguinis, 1985, 48:362–5.

22. Muller A et al. Mosaicisme Rh par mutation dans une gemellite monozygote. [Rh mosaicism by mutation in monozygotic twins.] Revue française de transfusion et immuno-hématologie, 1978, 21: 151–64.

23. Jenkins WJ, Marsh WL. Somatic mutation affecting the Rhesus and Duffy blood group systems. Transfusion, 1965, 20:6–10.

24. Northoff H et al. A patient mosaic for Rh and Fy antigens lacking other signs of chimerism or chromosomal disorder. Vox sanguinis, 1984, 47:164–9.

25. Avent ND. The rhesus blood group system: insights from recent advances in molecular biology. Transfusion medicine reviews, 1999, 13:245–66.

26. Avent ND, Martin PG. Kell typing by allele-specific PCR (ASP). British journal of haematology, 1996, 93:728–70.

27. Sambrook J, Fritsch E, Maniatis T. Molecular cloning: a laboratory manual, 2nd ed. New York, Cold Spring Harbor, 1989.

28. Wagner FF. Molecular basis of weak D phenotypes. Blood, 1999, 93:385–93.