Inoue T, Kira R, Nakao F, Ihara K, Bassuny WM, Kusuhara K, Nihei K, Takeshita K, Hara T. Contribution of the Interleukin 4 Gene to Susceptibility to Subacute Sclerosing Panencephalitis. Arch Neurol. 2002;59(5):822-827. doi:10.1001/archneur.59.5.822
Copyright 2002 American Medical Association. All Rights Reserved. Applicable FARS/DFARS Restrictions Apply to Government Use.2002
Although the exact pathogenesis of subacute sclerosing panencephalitis (SSPE) remains to be determined, both viral and host factors seem to be involved.
To identify host genetic factors involved in the development of SSPE.
We investigated the association of polymorphisms in the T helper (Th)1 and Th2 cytokine, and related genes (interferon [IFN]-γ, IFN-γ receptor 1 [IFN-γ R1], IFN-γR2 [IRF-1], interleukin 12 receptor β 1 [IL-12Rβ1], IL-4, IL-4R, and IL-10 genes) with SSPE in Japanese subjects.
A significant association (P = .03) was observed between SSPE and the T allele of the biallelic polymorphism at position −589 in the promoter region of the IL-4 gene. The IRF-1 allele 1 tended to interact with the IL-4 promoter −589 Tgenotype in the development of SSPE (P = .06), as judged on logistic regression analysis. The frequency of the genotype combination of IL-4 promoter −589 T and IRF-1 allele 1 (at least 1 allele) in patients with SSPE was much higher than that in the controls (47.7% vs 22.0%; P = .003, χ2 analysis). However, there was no association between other polymorphisms and SSPE.
To our knowledge, this study is the first to demonstrate the possibility that the IL-4 promoter gene –589 T gene polymorphism with increased IL-4 synthesis in combination with IRF-1 allele 1 confers host genetic susceptibility to SSPE in Japanese subjects.
SUBACUTE SCLEROSING panencephalitis (SSPE) is a slowly progressive central nervous system complication of measles. Although the exact pathogenesis of SSPE remains to be determined, both viral and host factors seem to be involved in it.1,2 As viral factors for the development of SSPE, measles viruses (MVs) isolated from the central nervous system of patients with SSPE are replication defective and have extensive mutations within the envelope-associated genes. However, matrix gene mutations thought to be characteristic of SSPE viruses have also been found in recent clinical isolates.3 Therefore, it is unclear whether these mutations are critical for the development of SSPE.
As a host factor, immaturity of the host immune system and central nervous system has been suggested to play a role in the increased risk for SSPE development among infants who have had measles before 2 years of age.1,4 Although no universal abnormality has been identified in the immune system of patients with SSPE, impairment of MV-specific cell-mediated immunity has been observed in a significant proportion,5,6 while anti-MV antibody production is enhanced in serum and cerebrospinal fluid. Thus, it is likely that the T helper (Th)1 function involved in cell-mediated immunity is generally down-regulated and the Th2 function implicated in antibody production is well preserved. In our previous study on live MV-specific Th1/Th2 cytokine production by peripheral blood mononuclear cells, most patients with SSPE exhibited decreased MV-specific Th1 cytokine production and preserved Th2 cytokine synthesis.7 Thus, it is possible that a defect of the Th1 response and the preserved Th2 response toward MV in most patients with SSPE might reflect the persistence of a relative dominance of the Th2 response over Th1 at the time of the initial measles infection.7 Enhanced Th2 polarization at the time of the initial measles infection may result in insufficient elimination of MV and a higher frequency of MV persistence. In the murine model of experimental MV encephalitis, MV-specific T cells from susceptible C3H mice produce smaller amounts of Th1 cytokines than those from resistant BALB/c mice.8
To determine the genetic background of the Th1/Th2 cytokine responses in patients with SSPE, we have mainly investigated functional or disease-associated polymorphisms of the Th1 and Th2 cytokine and related genes. The interleukin 4 (IL-4) promoter 589 C/T,9- 11 IL-4 receptor α chain (IL-4R) codon 50 Ile/Val,12IL-10 promoter 627 C/A,13 interferon-γ (IFN-γ) gene CA repeat, and IFN regulatory factor-1 (IRF-1) gene GT repeat14 polymorphisms are associated with Th2-shifted cytokine production or Th2-dominant disorders. In addition, the IFN-γ receptor 1 (IFNGR1) codon 14 Val/Met and IFNGR2 codon 64 Gln/Arg polymorphisms have also been examined, as their combination has been associated with systemic lupus erythematosus, a disorder of the Th1/Th2 balance.15 As Th1-inducing cytokine receptors, the IL-12 receptor β 1 chain (IL-12RB1) codon 214 Gln/Arg, 365 Met/Thr, and 378 Gly/Arg polymorphisms were included because of their association with mycobacterial infection, in the clearance of which Th1 cytokines play a major role.16 To our knowledge, this study is the first to demonstrate the possibility that the IL-4 promoter gene −589 T polymorphism with increased IL-4 synthesis in combination with IRF-1 allele 1 confers host genetic susceptibility to SSPE in Japanese subjects.
Thirty-eight Japanese patients with SSPE (25 male and 13 female subjects) and 100 healthy Japanese individuals composed the study population. All of the patients with SSPE fulfilled the diagnostic criteria, for example, clinical features, increased MV antibody titer in cerebrospinal fluid, and typical electroencephalography showing periodic slow-wave complexes early in the disease.1,4 The age of onset of SSPE in the study population ranged between 2 and 15 years (mean, 8.0 years). Thirty-three patients had natural measles occurring between the ages of 0.4 and 4 years (mean, 1.3 years), the measles history being unknown in the other 5 patients, including 1 with a history of live attenuated measles vaccination. The control subjects were randomly selected from among healthy school children. The control group was not age-matched for measles or measles vaccination. Informed consent was obtained from the subjects and/or their parents.
Genomic DNA was extracted from peripheral blood using a commercially available blood kit (QIAamp DNA Blood Kit; Qiagen, Tokyo, Japan). Each polymerase chain reaction (PCR) reaction was carried out with 20 ng of genomic DNA, 12.5 pmol of each primer, 0.6 U of Taq DNA polymerase (Promega, Madison, Wis), and 5 nmol of each deoxynucleoside triphosphate, in a total volume of 25 µL, using a PCR thermal cycler (Thermal Cycler MP; TaKaRashuzo Corp, Otsu, Japan). The sequences of the primers and probes used in this study are given in Table 1.
The C/T transition polymorphism at position –589 in the promoter region of the IL-4 gene was analyzed by PCR-restriction fragment length polymorphism (PCR-RFLP) as previously described.9 The PCR profile used was as follows: initial denaturation at 93°C for 5 minutes, followed by 36 cycles of 93°C for 1 minute, 48°C for 1 minute, and 72°C for 1 minute, with final extension at 72°C for 3 minutes. The PCR products were digested with AvaII at 37°C for 1 hour, followed by separation in 3% agarose gels.
The IL-4R codon 50 Ile/Val genotype analysis was performed using the allele-specific amplification method with a TaqMan fluorogenic probe (TaqMan-ASA).17 The Ile and Val alleles were detected with Ile- and Arg-specific primers, respectively, with a common primer. A one-base mismatch was introduced at the 3′ end of each ASA primer. The PCR amplification was carried out, in a total volume of 25 µL, with 2 sets of ASA primers in the presence of a TaqMan probe using a sequence detection system (PRISM 7700 Sequence Detection System; PE Biosystems, Foster City, Calif), which detects fluorescence emission from the reporter dye in each PCR cycle. The PCR conditions were as follows: initial denaturation at 95°C for 10 minutes, followed by 40 cycles of 95°C for 15 seconds, and 60°C for 1 minute. The fluorescence signal was monitored in real time to determine the threshold cycle at which the fluorescence emission level exceeded the baseline. The Ile or Val allele could be clearly distinguished based on the differences in the threshold cycles.
The genotypes for IL-10 promoter –627 C/A were determined by means of PCR–single-strand conformation polymorphism (PCR-SSCP). The PCR conditions were initial denaturation at 94°C for 5 minutes, followed by 35 cycles of 94°C for 30 seconds, 60°C for 30 seconds, and 72°C for 30 seconds, with final extension at 72°C for 5 minutes. The PCR products were mixed with the same volume of deionized formamide, denatured for 5 minutes at 95°C, and then run on GeneGel Excel 12.5/24 (Pharmacia Biotech, Uppsala, Sweden) at 25 mA and 15°C for 1½ hours. Subsequent silver staining revealed the variable mobilities of conformational fragments, which corresponded to the –627 C and A genotypes.
The genotypes for IL-12RB1 codons 214 Gln/Arg, 365 Met/Thr, and 378 Gly/Arg were defined by means of PCR-SSCP, TaqMan-ASA, and PCR–restriction fragment length polymorphism (RFLP), respectively. The PCR profile used for codons 214 Gln/Arg and 378 Gln/Arg was: 94°C for 30 seconds, 60°C for 30 seconds, and 72°C for 30 seconds, for 35 cycles, with final 5-minute extension at 72°C. Initial denaturation was conducted at 94°C for 5 minutes. Single-strand conformation polymorphism analysis was performed at 25 mA and 20°C for 75 minutes. For RFLP, the PCR products were digested with AvaII at 37°C for 1 hour. TaqMan-ASA was performed as described above. The Met and Thr alleles at codon 365 were detected with Met- and Thr-specific primers, respectively, with a common primer. A 1-base mismatch was introduced at the 3′ end of each allele-specific primer.
The IFNGR1 gene codon 14 Val/Met and IFNGR2 gene codon 64 Gln/Arg genotypes were defined by means of PCR-SSCP, as described previously.14 The PCR conditions for IFNGR1 were initial denaturation at 94°C for 5 minutes, followed by 40 cycles of 94°C for 30 seconds, 60°C for 30 seconds, and 72°C for 30 seconds, with final extension at 72°C for 7 minutes. The amplification conditions for IFNGR2 were the same except for an annealing temperature of 50°C. Single-strand conformation polymorphism analysis was performed at 25 mA and 20°C for 1½ hours, and 10 mA and 5°C for 3½ hours, respectively.
The region containing the CA repeat polymorphism within the first intron of the IFN-γ gene was amplified by PCR, as described previously.14 The 5′ end of the forward primer was fluorescently labeled with 6-carboxyfluorescein dye. The PCR conditions were as follows: initial denaturation at 95°C for 5 minutes, followed by 30 cycles of 95°C for 30 seconds, 56°C for 30 seconds, and 72°C for 1 minute, with final extension at 72°C for 5 minutes. Genotyping was performed in a mixture of amplified products and an internal size standard with a genetic analyzer (PRISM 310; PE Biosystems).
The GT repeat polymorphism of intron 7 in the IRF-1 gene was determined by PCR, as described previously.14 The 5′ end of the forward primer was fluorescently labeled with hexachloro-6-carboxyfluorescein dye. The PCR conditions were initial denaturation at 95°C for 5 minutes, followed by 30 cycles of 95°C for 30 seconds, 56°C for 30 seconds, and 72°C for 1 minute, with final extension at 72°C for 5 minutes. Genotyping was performed as described above.
Differences between allele or genotype frequencies in the 2 groups were evaluated by means of the χ2 analysis with a 2 × 2 or contingency table. Whole allele distributions were analyzed by means of the χ2 test with a 2 × 7 contingency table. When at least 1 cell number was not more than 5, the 2-sided Fisher exact test was used for the χ2 value. Logistic regression analysis was performed to identify any interaction among the polymorphisms. P<.05 was considered to be statistically significant.
The frequencies of each allele in the 38 patients with SSPE and 100 healthy controls are given in Table 2 and Table 3. A significant difference in the allele frequencies between patients and controls was found for the IL-4 promoter –589 C/T gene polymorphism. The frequency (0.789) of the T allele in patients with SSPE was significantly higher than that (0.655) in healthy controls (P = .03).
There were no significant differences in the allele frequencies of the IL-4R 50 Ile/Val, IL-10 promoter –627 C/A, IL-12RB1214 Gln/Arg, IL-12RB1 365 Met/Thr, IL-12RB1378 Gly/Arg, IFNGR1 14 Val/Met, IFNGR2 64 Gln/Arg, IFN-γ CA repeat, and IRF-1 GT repeat polymorphisms.
As the IL-4 promoter –589 C/T gene polymorphism was associated with SSPE, interactions between this polymorphism and the other 9 polymorphisms were evaluated by logistic regression analysis. IRF-1 allele 1 tended to interact with the TT genotype in the development of SSPE (P = .06). The frequency of the genotype combination of IL-4 promoter –589 T and IRF-1 allele 1 (at least 1 allele) in patients with SSPE was much higher than that in controls (47.7% vs 22.0%; P = .003, χ2 analysis). Among 5 patients with SSPE older than 2 years (ie, 4, 2, 4, 3, and 3 years) at the time of the initial MV infection, 4 were homozygous for the IL-4–589 T allele, and the other was heterozygous. The clinical characteristics of each individual patient with SSPE along with his or her IL-4 promoter –589 C/Tgenotype are shown in Table 4.
For this study, samples from 13 of the 15 patients with SSPE with low or no MV-specific IFN-γ production in our previous study7 were available. No correlation was observed between MV-specific IFN-γ production or disease progression and gene polymorphisms. In addition, MV-specific IL-4 production levels were below the detection limit in all patients with SSPE except 1 in the previous study.
To our knowledge, in this study, we have first demonstrated that the IL-4–589T allele is significantly associated with SSPE. In addition, the association with SSPE was more significant for the genotype combination of IL-4 promoter –589 T and IRF-1 allele 1. Thus, it is possible that polymorphisms of the IL-4 promoter and IRF-1, as host genetic factors, contribute to a predisposition to SSPE in Japanese subjects.
The Th1 and Th1-inducing cytokines such as IL-2, IL-12, IFN-γ, and tumor necrosis factor α preferentially induce cell-mediated immunity, while Th2 cytokines such as IL-4, IL-6, and IL-10 primarily support antibody production. Interleukin 4 is a key cytokine for the Th2 response, which induces activation and maturation of B cells as well as differentiation of immature Th cells into Th2 cells, while IL-4 inhibits the differentiation and function of the Th1 phenotype, especially IFN-γ production. In its promoter region, the IL-4 –589T allele is associated with increased IL-4 gene promoter activity,9- 11,18 leading to a relative dominance of the Th2 response. Actually, the genotype combination of IL-4 promoter –589 T and IRF-1 allele 1 might further promote Th2 polarization, based on our recent observation that this combination is significantly associated with atopic asthma, a Th2-dominant disorder.14 In addition, the IL-4–589 T allele has been suggested to play a role in the interaction between the host and virus.18
Measle virus infection itself influences concurrent and subsequent Th1 vs Th2 immune responses. Measles is associated with Th1 activation (increased IFN-γ and IL-2 levels) before and during the rash phase, followed by preferential Th2 activation (decreased IFN-γ and IL-2 levels, and increased IL-4 level) during the convalescent phase.19,20 The Th1 response seems to be crucial for clearance of MV from the blood and other tissues mostly within the first 1 to 2 weeks after the onset of the rash. However, the Th2 response leading to anti-MV antibody production may cause decreased recognition of infected cells by the immune system through the reduction of viral antigen expression,21,22 and thereby result in the establishment of persistent MV infection. Therefore, it is possible that the genetic predisposition to Th2 dominance in patients with SSPE also plays a role in the persistence of MV and the development of SSPE through reduction of the Th1 response and enhancement of anti-MV antibody production.
In our previous study on 15 patients with SSPE, we showed that low or no MV-specific IFN-γ production was associated with rapid disease progression.7 Determination of the cytokine-related gene polymorphisms in 13 of the 15 patients revealed that there was no correlation between MV-specific IFN-γ production or disease progression and gene polymorphisms. Once SSPE has developed, nongenetic and/or other genetic factors may play major roles in the decreased MV-specific IFN-γ production and disease progression.
An immature immune system at the time of the initial measles infection is considered to play a role since MV infection in early infancy leads to development of SSPE at a higher frequency than when measles occurs later in life, as in the case of persistent hepatitis B virus infection.23 Neonatal tolerance is thought to play a role in the establishment of chronic hepatitis B virus infection in children born to hepatitis B virus–infected mothers. In animals with an immature immune system, selection of either the tolerance or protective immunity pathway is determined in part by the dose and form of antigen presented.24- 26 Such tolerance is not simply the result of immunologic immaturity but rather is correlated with the induction of a nonprotective Th2 cytokine response. Our patients with SSPE older than 1 year at the time of the initial MV infection showed a higher frequency of the IL-4–589 T allele than those 1 year old or younger at the time of measles infection, although the sample size was too small for statistical analysis. Thus, Th2 polarization during and after MV infection might result in insufficient elimination and persistence of MV in cases with an immature immune system or Th2-shifted cytokine gene polymorphisms. Host genetic factors for SSPE might be difficult to identify in such cases, as environmental factors contributing to early MV infection greatly influence the risk of SSPE.27,28 Further investigation of host genetic factors for SSPE is necessary in other countries with different environmental and genetic backgrounds.
Accepted for publication January 8, 2002.
Author contributions: Study concept and design (Drs Inuoe, Kira, Ihara, Kusuhara, Takeshita, and Hara); acquisition of data (Drs Inuoe, Kira, Nakao, Bassuny, Kusuhara, Nihei, and Hara); analysis and interpretation of data (Drs Inuoe, Kira, Ihara, Ihara, Kusuhara, and Hara); drafting of the manuscript (Drs Inuoe, Kira, Kusuhara, and Hara); critical revision of the manuscript for important intellectual content (Drs Kira, Nakao, Ihara, Bassuny, Kusuhara, Nihei, Takeshita, and Hara); obtained funding (Drs Kira, Ihara, Kusuhara, Nihei, Takeshita, and Hara); administrative, technical, and material support (Drs Inuoe, Kira, Nakao, Ihara, Bassuny, Kusuhara, Takeshita, and Hara); study supervision (Drs Kira, Ihara, Kusuhara, Takeshita, and Hara).
This study was supported in part by grants from the Ministry of Health and Welfare of Japan, and the Ministry of Education, Science, Sports and Culture of Japan, Tokyo.
We thank H. Hattori, MD (Osaka City University Medical School), S. Yamashita, MD (Kanagawa Children's Medical Center), N. Koide, MD (National Iwaki Hospital), H. Aiba, MD (Shizuoka Children's Hospital), T. Okada, MD (Kochi Medical School), F. Hamada, MD (Hosogi Hospital), N. Koyama, MD (Toyohashi Municipal Hospital), Y. Hirata, MD (Hamamatsu Medical Center), C. Baba, MD (Red Cross Nagasaki Atomic Bomb Hospital), A. Ono, MD (Saiseikai Izumio Hospital), A. Tomoda, MD (Kumamoto University), M. Funahashi, MD (Tokyo Children's Rehabilitation Hospital), T. Kurokawa, MD (National Nishi-Beppu Hospital), R. Sakuta, MD (Dokkyo University Koshigaya Hospital), M. Miyazaki, MD (Tokushima University), K. Shioya, MD (National Nichinan Hospital), N. Nagano, MD (Asahikawa City Hospital), T. Ishizu, MD (National Saishunso Hospital), K. Gondo, MD and Y. Tokunaga, MD (Kyushu University), and K. Watanabe, MD (Kagoshima Municipal Hospital) for providing us with samples from their patients, as well as Dr N. Kinukawa, PhD (Kyushu University) for the advice on statistical analyses.
Corresponding author and reprints: Ryutaro Kira, MD, PhD, Department of Pediatrics, Graduate School of Medical Sciences, Kyushu University, 3-1-1 Maidashi, Higashi-ku, Fukuoka 812-8582, Japan (e-mail: email@example.com).