OUP user menu

Comprehensive and Efficient HBB Mutation Analysis for Detection of β-Hemoglobinopathies in a Pan-Ethnic Population

Owen T. M. Chan MD, PhD, Kenneth D. Westover MD, PhD, Lisa Dietz, James L. Zehnder MD, Iris Schrijver MD
DOI: http://dx.doi.org/10.1309/AJCP7HQ2KWGHECIO 700-707 First published online: 1 May 2010


Current methods that assay hemoglobin β-globin chain variants can have limited clinical sensitivity when applied techniques identify only a predefined panel of mutations. Even sequence-based assays may be limited depending on which gene regions are investigated. We sought to develop a clinically practical yet inclusive molecular assay to identify β-globin mutations in multicultural populations. We highlight the β-globin mutation detection assay (β-GMDA), an extensive gene sequencing assay. The polymerase chain reaction (PCR) primers are located to encompass virtually all hemoglobin β locus (HBB) mutations. In addition, this assay is able to detect, by gap PCR, a common large deletion (Δ619 base pair), which would be missed by sequencing alone. We describe our 5-year experience with the β-GMDA and indicate its capability for detecting homozygous, heterozygous, and compound heterozygous sequence changes, including previously unknown HBB variants. The β-GMDA offers superior sensitivity and ease of use with comprehensive detection of HBB mutations that result in β-globin chain variants.

Key Words:
  • Thalassemia
  • β-Globin
  • Ethnicity
  • Hemoglobinopathy
  • Molecular diagnosis

Numerous disorders of the β-globin chain of hemoglobin lead to different disease phenotypes.1,2 Of these, β-thalassemia is a subset of the β-hemoglobinopathies characterized by a hereditary anemia with a wide phenotypic spectrum that can have significant morbidity and mortality.2,3 This autosomal recessive condition afflicts thousands of people of many ethnicities worldwide.4 In India, the carrier frequency of β-thalassemia spans 0.3% to 15%, depending on the ethnic subgroup.5 On the African continent, the β-thalassemia carrier frequency varies from 1% to 13%.5 In Europe, the percentage of the population carrying a significant variant associated with β-thalassemia ranges from 0.12% in Finland to 15% in Cyprus.6 In the United States, screening in California identified 1 newborn with β-thalassemia for every 55,000 births.7 Exact carrier frequencies for the multiethnic population of the United States overall have, to our knowledge, not been reported. During the next 20 years, it is predicted that more than 900,000 children will be born with clinically significant β-thalassemia or hemoglobin E-β-thalassemia.8

The β-thalassemias exhibit a range of severities, each corresponding to an absence or reduction of β-globin protein synthesis. These β-thalassemia phenotypes are related to the myriad mutations that affect the β-globin gene (HBB) on chromosome 11p15.5 (OMIM number +141900), and different populations have their own mutation spectrum.9 More than 730 variants of the β-globin chain have been characterized, and more than 200 of these account for mutations causing β-thalassemia.10 The sequence variants range from point mutations to large deletions and affect HBB gene transcription, messenger RNA processing, translation, or gene product structure.2 While the majority of the variants are rare, 21 mutations account for more than 80% of all β-thalassemia determinants Table 1.2,11

A variety of assays have been developed to detect β-thalassemia. To screen for mutations, methods include denaturing gradient gel electrophoresis,12,13 temporal temperature gel electrophoresis,14 mismatch analysis,15,16 single-strand conformation polymorphism electrophoresis,17,18 and heteroduplex analysis with denaturing high-performance liquid chromatography.1923 In addition, specific detection strategies can be used for known mutations, such as techniques that incorporate allele-specific oligonucleotide probes,2426 the amplification refractory mutation system,2730 allele-specific polymerase chain reaction (PCR),31 reverse dot blot,32,33 restriction fragment length polymorphisms,34 single base extension,35,36 PCR–enzyme-linked immunosorbent assay,37 real-time PCR,38 and array-based technologies.3944 Mass spectrometry has also been used to identify hemoglobin variations on the nucleic acid and protein levels.45,46

Despite these numerous methods to assay for β-thalassemia, mutations can still go undetected, especially when using detection strategies that identify only a specific subset of the various β-globin variants. Direct DNA sequencing enables more comprehensive detection of known and unknown β-globin gene mutations.4751 Sequencing allows the identification of most β-hemoglobinopathies, but the location of the primers is critical because mutations in the HBB gene are not limited to the exons and their direct splice sites. Sequence analysis cannot identify mutations caused by large deletions (or insertions) of 1 or more exons because primers would not be able to bind to a deleted sequence. Thus, the resulting sequence tracing would show only the (hemizygous) sequence from the present allele. This can be remedied by using a technique such as multiplex ligation-dependent probe amplification as a follow-up test, if so desired.52,53

Although heritable large duplications or deletions in the β-globin gene are typically rare, the 619-base-pair (bp) deletion mutation (Δ619 bp), which encompasses exon 3, is an exception that has greater prevalence.5456 This deletion, associated primarily with certain Indian and Pakistani subpopulations,57 comprises up to 7.5% of the β-thalassemia subtypes found in the Indian population.58,59 To screen β-thalassemia mutations in the Southeast Asian and Indian populations, Wang et al60 designed a limited sequencing assay that also could detect the Δ619 bp allele. These investigators combined gap PCR (an amplification reaction for which primers flank the deletion site) to recognize this deletion mutation with multiplex minisequencing, which used primers to recognize 15 additional common mutations in their targeted population. Their combined strategy allowed the detection of this large deletion mutation and, in conjunction with minisequencing, enabled detection of more than 90% of β-thalassemia alleles in Southeast Asia and India. In the United States, however, the ethnic diversity is considerable. We hypothesized that a more inclusive mutation detection approach could be beneficial.

View this table:
Table 1

In the present article, we describe a combined gap PCR and broad sequencing method that combines the high sensitivity of sequencing with the ability to detect the Δ619 bp allele. As a result, this assay is able to identify all of the most prevalent β-thalassemia mutations mentioned earlier,2,11 in addition to detecting almost all other β-globin variants. We report our 5-year experience with this method.

Materials and Methods

Blood Specimens and Genomic DNA Extraction

Patient blood samples were submitted in EDTA, acid-citrate-dextrose, or sodium citrate. To extract DNA, the specimens were processed by using Qiagen minicolumns or the Qiagen EZ1 extraction procedure (Qiagen, Valencia, CA).

Multiplex Minisequencing Screen

The strategy to detect 15 β-thalassemia mutations with a multiplex minisequencing assay was described by Wang et al.60 This protocol was slightly modified, where the thermocycling conditions consisted of 1 denaturing cycle at 95°C for 10 minutes followed by 35 cycles of denaturing at 95°C for 30 seconds, annealing at 58.5°C for 1 minute, and extension at 72°C for 2 minutes. Final extension was at 72°C for 10 minutes.

Genomic DNA Amplification and Full Sequencing

To amplify and sequence the β-globin gene via PCR, primers Table 2 were designed and subsequently synthesized at Operon (Huntsville, AL). First, genomic DNA was amplified in 2 different PCR master mixes (PCR I and II), each containing a total volume of 50 μL per reaction. PCR I contained ddH2O (Teknova, Hollister, CA), 5 U of TaqGold (Applied Biosystems, Foster City, CA), PCR amplification buffer (Applied Biosystems), a final concentration of 1 mmol/L magnesium chloride (Applied Biosystems), 150 μmol/L 4dNTP mix (Pharmacia, Piscataway, NJ), 20 pmol of primer A, 20 pmol of primer B, 20 pmol of primer D, and 100 to 200 ng of DNA. PCR II contained the same reagents except for the primers, which were 20 pmol of primer C and 20 pmol of primer D. The thermocycling conditions were 1 denaturing cycle at 95°C for 10 minutes followed by 35 cycles of denaturing at 95°C for 30 seconds, annealing at 58.5°C for 1 minute, and extension at 72°C for 2 minutes. Final extension was at 72°C for 10 minutes. The amplification products were then electrophoresed in a 1% agarose gel to assess PCR efficacy and product size. The presence of the 619-bp deletion is assessed on this gel.

Before sequencing, the amplicons were purified with the QIAquick PCR Purification Kit (Qiagen) or with ExoSAP-IT (USB, Cleveland, OH). After purification, 5 μL of the amplicons served as templates for sequencing using the ABI PRISM BigDye Terminator version 3.1 Cycle Sequencing Kit (Applied Biosystems). The primers used for sequencing (PCR I, A and E; PCR II, C and F) were used at a final concentration of 1 μmol/L. The thermocycling conditions consisted of 25 cycles of denaturing at 96°C for 10 seconds, annealing at 50°C for 5 seconds, and extension at 60°C for 4 minutes. The resultant sequencing amplicons were purified with a 2.2% sodium dodecyl sulfate solution in Centri-Sep spin columns (Princeton Separations, Adelphia, NJ) and electrophoresed in an ABI PRISM 3130xl Genetic Analyzer (Applied Biosystems). The sequencing data were analyzed with Sequencing Analysis Software, version 5.2 (Applied Biosystems) or Mutation Surveyor (Softgenetics, State College, PA).

View this table:
Table 2

β-Globin Mutation Detection Assay

Figure 1 depicts the human β-globin gene with corresponding β-thalassemia mutations. To detect these mutations, gap PCR and direct sequencing were combined into a 2-phase assay. The primers used in the assay are listed in Table 2 and also graphically illustrated in Figure 1. As the first phase, the gap PCR, characterized by Wang et al,60 was conducted to detect the Δ619 bp mutation. Two PCR reactions (PCR I and II) were performed on genomic DNA. An unaffected wild-type patient would yield a 1,457-bp product from PCR I (primer pair A and B) and a 1,212-bp product from PCR II (primer pair C and D). In an affected person with a homozygous Δ619 bp allele, PCR I would yield a longer 1,671-bp product from the A and D primer pair (due to the deletion of the B primer binding site), and PCR II would yield a truncated product of 593 bp from the C and D primer pair. Heterozygous persons would have a combination of the 1,457- and 1,671-bp products from PCR I and the 1,212- and 593-bp products from PCR II.

Figure 1

Genomic map of the HBB gene depicting β-thalassemia mutations. The figure was adapted from the UCSC (University of California Santa Cruz) Genome Bioinformatics (http://genome.ucsc.edu) human March 2006 assembly using data from the HbVar database (a relational database of human hemoglobin variants and thalassemia mutations) on the globin gene server (http://globin.bx.psu.edu/hbvar). Genome base positions of chromosome 11 are indicated on the scale at the top of the figure. The exons are represented as black boxes with intervening sequences (IVS). The known β-thalassemia mutations and their corresponding gene locations and representative sizes are depicted graphically as bars above the HBB gene. The Δ619-base-pair mutation (base positions 5,203,195 to 5,203,813) is listed above the region spanning part of IVS II and all of exon 3. The scale of the figure does not allow individual discrimination of the various small sequence variants. The polymerase chain reaction primers used in the amplification and sequencing assay are illustrated as gray arrows (not drawn to scale) indicating their forward or reverse direction. Base positions of sequencing primers: A = 5,204,985-5,205,004; E = 5,204,309-5,204,328; C = 5,203,905-5,203,926; F = 5,203,208-5,203,227. HBB, hemoglobin β locus.

The second phase in our assay used dye termination sequencing of the resultant PCR I and II amplicons. The sequencing primers (PCR I, A and E; PCR II, C and F) are listed in Table 2 and graphically depicted in Figure 1. The reverse sequencing product from PCR I is 696 bp and spans exon 1, intervening sequence (IVS) I, and exon 2. PCR II’s reverse sequencing product is 719 bp long and covers a portion of IVS II and all of exon 3. Combined together, the forward and reverse sequencing data from PCR I and PCR II encompass virtually all of the known point mutations and small deletions of β-thalassemia (Figure 1).


Summary of β-Hemoglobinopathies

The β-globin mutation detection assay (β-GMDA) was implemented in our laboratory in March 2004 and continues to be performed to the present. The multiplex minisequencing assay described by Wang et al60 was used in our laboratory from December 2003 to February 2006. During the period when both techniques were offered for diagnosis, physicians were given the option to order either assay or to order the sequencing assay following the multiplex minisequencing assay if the latter was not definitive, such as would be the case when only 1 mutation was identified in an affected patient.

In Table 3, we summarize the β-hemoglobin variant data from 201 patients during a 5-year period (December 2003 to December 2008). Although RBC indices and ethnicity are requested by our laboratory, for most of our patients, this information is not provided and, unfortunately, could not be included in this study. The various mutations detected by the 2 assay systems were compiled with their respective frequencies and percentages of total identified alleles (n = 61). A total of 28 different β-globin alleles were identified. Of these, 19 were β-thalassemia mutations and 8 were β-hemoglobin variants associated with other conditions, such as hemoglobin C and hemoglobin S. The β-GMDA also identified just 1, to our knowledge, previously unreported variant, HBB:c.246C>A (Leu82Leu), which was thought to be most likely clinically insignificant because it was a silent mutation.

View this table:
Table 3

β-Globin Mutation Detection Assay

The β-GMDA is not limited to the identification of specific mutations; rather, known and unknown changes in the HBB gene can be detected by this sequencing-based method. With this assay, the HBB:c.316-90A>G variant in IVS II was discovered by our laboratory, and it was reported subsequently to the HbVar database on the globin gene server (IVS-II-761 A>G; http://globin.bx.psu.edu/hbvar). Of the 201 patients, 58 (28.9%) had detectable mutations. Three of these patients were compound heterozygous. Two patients were identified by the assay as being homozygous for HBB:c.47G>A (Trp16Stop) and HBB:c.79G>A (hemoglobin E), respectively. Two other patients in our tested population were compound heterozygous (patient 1: HBB:c.93-21G>A [IVS I] and HBB:c.114G>A [Trp38Stop]; and patient 2: HBB:c.79G>A [hemoglobin E] and HBB:c.441_442insAC [hemoglobin Tak]). Thus, in addition to heterozygous changes, the β-GMDA is capable of detecting homozygous and compound heterozygous gene alterations.


Elucidationofthepathophysiologyofthalassemiaspanned several decades. The imbalance of α or β globin chains was realized in the 1960s as globin synthesis was measured.61,62 During the 1970s and 1980s, investigators began to identify the molecular origins of the thalassemia conditions.56,6365 With the advancement of molecular genetic techniques, most of the globin chain gene variants were understood and definitively characterized by the late 1990s.66,67

β-thalassemia mutations are composed primarily of mis-sense mutations or other small sequence changes.2 In contrast to α-thalassemia, a very small subset of the mutations is composed of large deletions. Only the Δ619 bp mutation, which is most prominently observed in some subpopulations from India and Pakistan,5759,68 has increased prevalence. While sequencing strategies do not typically detect large deletions,49,50 the β-GMDA was designed to detect the Δ619 bp allele in the β-globin gene by combining the sequencing approach with a preceding gap PCR step.60 Fourteen other large deletions that range from 290 bp to greater than 60 kb2 would not be detected by the β-GMDA. Of the large β-globin deletions, however, only Δ619 bp is relatively common, and the remaining are extremely rare.2 If a large deletion mutation other than Δ619 bp is present in homozygous form, the primer binding sites will be absent, and no PCR product will be generated using the β-GMDA. Therefore, a negative amplification phase would suggest a technical amplification failure or the presence of homozygous, large, non–619-bp deletion mutations, which are exceedingly rare.

Panel testing for β-thalassemia typically recognizes only a limited number of mutations and is designed to find selected mutations among specific ethnic groups.24,27,37,60,69 A more comprehensive technique that identifies mutations from multiple ethnicities is expected to have greater sensitivity and, thus, greater usefulness in today’s multicultural US society. Our laboratory used the multiplex minisequencing screen (MMS) described by Wang et al60 for almost 2 years concurrently with the β-GMDA. Both methods were characterized by excellent performance. The MMS has primer pairs that recognize 15 β-globin mutations. Table 3 lists the total cumulative alleles and their frequencies identified by the 2 methods during a 5-year period. As indicated in Table 3, 28 different β-globin variants were detected.

Compared with the β-GMDA, the MMS, not surprisingly, has limited sensitivity because it can only identify 15 β-globin variants in addition to the Δ619 bp mutation. As shown in Table 3, the MMS primer panel enables the detection of only 12 of these 28 variants. In addition, samples from 4 persons of the 201 patient sample population were assayed by the MMS and by the β-GMDA. These 4 people were initially screened as negative by the MMS. However, the β-GMDA subsequently revealed β-globin mutations (Trp16Stop, Trp38Stop, HBB:c.93-21G>A, and His118Arg) not detectable by the MMS primer panel. Together, these findings supported the need, for our patient population, to offer a comprehensive assay not limited by a specific mutation panel.

Our detection strategy was designed to be comprehensive in identification of numerous short-sequence alterations that dominate the expansive list of β-thalassemia mutations. In addition, the gap PCR component of our assay detects the Δ619 bp allele.60 This is a component that further increases the mutation detection capability.49,50 Because the majority of β-hemoglobin gene variants not associated with β-thalassemia (eg, hemoglobin S and hemoglobin C) also lie within the areas bounded by the sequencing primers used in the β-GMDA, our assay can be used for identification or confirmation of other HBB-associated diseases or the carrier status of such, as well. Although the assay does not cover the complete IVSs, it encompasses all exonic and intronic regions in which clinically significant changes have been reported (Figure 1). Thus, our detection system recognizes almost all β-thalassemia mutations and other β-hemoglobin variants (Table 3).

High-performance liquid chromatography and protein electrophoresis are routine methods often used to screen for β-hemoglobinopathies and may be sufficient for most clinical situations. However, there are circumstances when the β-GMDA would provide valuable diagnostic information not gleaned by conventional assays. For example, the molecular analysis of the β-GMDA can assess the β-hemoglobinopathy carrier state of a patient. This has important clinical implications, especially for couples planning to have children. In addition, the β-GMDA is amenable to prenatal diagnosis through the analysis of fetal DNA samples. Other clinical applications include providing prognostic data for patients affected with β-hemoglobinopathies. Especially for β-thalassemia with its numerous variants and variable severities,67 the β-GMDA identifies a patient’s specific β-globin mutation and, in turn, enables a physician to provide the appropriate treatment and counseling. Therefore, the β-GMDA provides better discriminatory power over routine screening methods and could be used as clinically needed.

In our patient population during a 5-year span (Table 3), 46% (13/28) of the β-thalassemia variants detected were among the 21 most prevalent β-thalassemia determinants worldwide (Table 1). The remaining 54% (15/28) of the variants, comprising 43% (26/61) of the identified alleles in our sample population, indicated that a significant subset of our ethnically diverse patients carried rarer β-thalassemia mutations. Also, as indicated in Table 3, the mutation panel–based MMS would not detect 57% (16/28) of the total detected β-globin variants. Therefore, this finding reaffirms the need to have an inclusive, comprehensive, non–mutation panel–based assay to detect the less prevalent mutations in our geographic region.

Automated DNA sequencing is a reliable, sensitive, and specific method that has become a routine tool in many laboratories and is in use for HBB testing in several clinical laboratories (http://www.ncbi.nlm.nih.gov/sites/GeneTests/?db=GeneTests). The β-GMDA capitalizes on the advantages that DNA sequencing offers over nonsequencing methods. In addition, it provides sequences that are not limited to exons and their exonintron boundaries, but rather includes virtually all areas of the gene where mutations have been described. The technology is advancing, leading to decreased processing times and increased ease of use.70 As the sequencing techniques mature, it is expected that the β-GMDA could be adapted to the various platforms that emerge.

The β-GMDA is a comprehensive and robust method for the detection of known and novel β-globin variants. The assay’s ability to detect virtually all of the small β-globin variations through inclusive sequencing and detection of the Δ619 bp via gap PCR confers high sensitivity to the technique and will be highly valuable for β-hemoglobinopathy detection in multicultural populations.


We thank Linda Gojenola for help and technical expertise.


  1. 1.
  2. 2.
  3. 3.
  4. 4.
  5. 5.
  6. 6.
  7. 7.
  8. 8.
  9. 9.
  10. 10.
  11. 11.
  12. 12.
  13. 13.
  14. 14.
  15. 15.
  16. 16.
  17. 17.
  18. 18.
  19. 19.
  20. 20.
  21. 21.
  22. 22.
  23. 23.
  24. 24.
  25. 25.
  26. 26.
  27. 27.
  28. 28.
  29. 29.
  30. 30.
  31. 31.
  32. 32.
  33. 33.
  34. 34.
  35. 35.
  36. 36.
  37. 37.
  38. 38.
  39. 39.
  40. 40.
  41. 41.
  42. 42.
  43. 43.
  44. 44.
  45. 45.
  46. 46.
  47. 47.
  48. 48.
  49. 49.
  50. 50.
  51. 51.
  52. 52.
  53. 53.
  54. 54.
  55. 55.
  56. 56.
  57. 57.
  58. 58.
  59. 59.
  60. 60.
  61. 61.
  62. 62.
  63. 63.
  64. 64.
  65. 65.
  66. 66.
  67. 67.
  68. 68.
  69. 69.
  70. 70.
View Abstract