Fetal microchimerism refers to a state in which cells from two or more genetically different individuals are present in the maternal body as a result of fetomaternal transfusion during pregnancy. Testing for fetomaternal microchimerism is based on the fact that the mother should carry only the X chromosome, and the test uses polymerase chain reaction (PCR) targeting the Y chromosome sequence that, barring blood transfusion or tissue transplantation, could come from a male fetus.
Temporary microchimerism occurs during pregnancy when fetomaternal transfusion of blood cells occurs across the placenta.1 After delivery, when the pregnancy has been terminated and the placenta has been expelled, fetal-origin cells should theoretically disappear from the maternal circulation. However, studies of women who have given birth to a male child have shown a high incidence of male cells in the blood, and these have been thought to have derived from the fetus.2 Moreover, some women have displayed male cells in the blood even several years after having given birth to a male child.2,3 This latter finding can be interpreted as meaning that fetal cells transferred from the fetus to the mother as a result of fetomaternal transfusion during pregnancy, then established permanent microchimerism in the maternal body. Permanent microchimerism was reportedly found in 30–50% of women who had given birth to a male child.3,4 In addition, although in theory it is impossible for the Y chromosome to be present in a woman who has never given birth, cells of male fetal origin have been detected at an incidence of 10–17% in such women.3,5 Conversely, Nelson et al6 reported a high rate of detecting male DNA in the peripheral blood of women with scleroderma who had given birth to a male child. Moreover, others have reported associations between fetomaternal microchimerism and autoimmune diseases such as scleroderma and lupus erythematosus.7,8
Microchimerism caused by fetal-origin cells has been detected in both healthy women and women with disease, but the clinical significance of this phenomenon remains unclear. In addition, little is known about the establishment of microchimerism in the case of spontaneous miscarriage or induced abortion in early pregnancy. One hypothesis holds that fetal-origin cells are immature during early pregnancy, and thus easily become implanted after curettage of the uterus.9 In light of this background, we estimated the incidence of persistent microchimerism after induced or spontaneous abortion in early pregnancy.
MATERIALS AND METHODS
With the approval of the Ethics Committee of Fukushima Medical University, written informed consent was obtained from each of the participating participants before enrolment in the study. Healthy women without hypertension, diabetes, or taking medication were enrolled on the basis of never having received a blood transfusion or had an abortion or male delivery. In theory, these women thus should not have carried any Y chromosomal material. From that population, a total of 100 women were registered after having undergone dilatation and curettage (D&C) for pregnancy termination or after spontaneous miscarriage during the period from February 2006 to October 2007 at any one of three obstetric institutions in Fukushima City.
Gestational age was determined by transvaginal ultrasonography. In the case of women who underwent induced abortion, crown-rump length of the fetus was measured and gestational age was decided. For women who experienced spontaneous miscarriage, when there was a fetus but a heart beat could no longer be found, a diagnosis of missed abortion was made, and gestational age was then decided from the crown-rump length. In the case of blighted ovum (ie, missed abortion in which no fetus was present), gestational age was calculated from last menses.
After the participant had provided written informed consent, 2 mL of peripheral blood was drawn into an ethylenediaminetetraacetic acid–treated test tube at each of three time points: before the abortion procedure (day 0), 7 days after abortion (day 7), and 30 days after abortion (day 30). These blood samples were stored at 4°C until testing. In addition, chorion obtained as a result of D&C was rinsed in tap water, and a portion was cut into 0.5–1.0-cm pieces that were stored frozen at –20°C.
After blood samples were centrifuged, 500 microliters of the buffy coat layer was dispensed into a tube containing red cell lying buffer (Genome Science Laboratories, Nagoya, Japan). After thorough mixing, the tube was centrifuged, the supernatant was discarded, and the pellet was twice more subjected to the same rinse procedure. The final pellet of white blood cells was suspended in 180 microliters of phosphate buffered saline buffer.
DNA was extracted from white blood cells using a QIAamp DNA Mini Kit (QIAGEN GmbH, Hilden, Germany). DNA concentration was adjusted to 100 micrograms/mL. Then, using AmpliTaq Gold DNA polymerase (Applied Biosystems, Foster City, CA), template DNA was subjected to nucleic acid amplification using the nested PCR method targeting the Y chromosome–specific DYS14 sequence.
The following four kinds of primers were used according to the methods of Lo et al10 Y1.5: DYS14–693F (5′-CTA GAC CGC AGA GGC GCC AT-3′), Y1.6: DYS14–931R (5′-TAG TAC CCA CGC CTG CTC CGG-3′) for first PCR, and Y1.7: DYS14–713F (5′-CAT CCA GAG CGT CCC TGG CTT-3′), Y1.8: DYS14–910R (5′-CTT TCC ACA GCC ACA TTT GTC-3′) for the second PCR. First PCR amplification was performed in a total volume of 50 microliters containing 0.4 micrograms of template DNA. The thermal cycling was as follows: denaturation at 95°C for 10 minutes, followed by 35 cycles of 94°C for 30 seconds, 57°C for 30 seconds, and final incubation at 72°C for 10 minutes. Second PCR amplification used 0.5 microliters of first product, and 25 cycles of thermal amplifications were done.
Electrophoresis of the reacted solution was performed in agarose gel, and the presence of fetal-origin male cells was determined on the basis of a band representing the DYS14 sequence. The sensitivity of PCR detection was 10−5, with 100% sensitivity for one male cell in 100,000 female cells. The negative control was Milli-Q water, which was negative for both the first and second PCR runs, whereas positive controls comprised DNA extracted from the peripheral blood of an adult male and DNA extracted after dilution of peripheral blood from an adult male to 1/105 with peripheral blood from an adult female. Nested PCR was performed twice per sample, in duplicate, and the sample was judged to be positive for sex-determining region of Y (SRY) if the DYS band was seen in even one of the four reaction tubes.
A portion of frozen and thawed chorion specimens was rinsed with phosphate buffered saline buffer, followed by pulverization using a pulverizer (Puluvei S-120, Sanki Kagaku, Tokyo, Japan) that had been chilled to –40°C. DNA was then extracted using a QIAamp DNA Mini Kit. Polymerase chain reaction using Y1.7/Y1.8 primer and electrophoresis were performed, and the sex of the fetus was determined on the basis of the presence or absence of a band of DYS14 sequence.
To avoid contamination of samples with Y chromosome, all procedures from collection of samples through to PCR were performed by female nurses/doctors/technicians, and the equipment used for each sample collection was disinfected with 0.5% hypochlorous acid.
Data were statistically analyzed using SPSS 11.5 software (SPSS Inc, Tokyo, Japan). A table was prepared showing mean±standard deviation for maternal age and gestational age, and the two participant groups were tested for significant differences using the Wilcoxon rank sum test. Data on the positive rate for microchimerism were analyzed using the Fisher exact test. Values of P<.05 were considered statistically significant.
Samples were given at day 0 by 100 participants, through to day 7 by 91 of the 100 participants, and through to day 30 by 76 participants. Analysis of chorion specimens showed that 44 of the 100 fetuses (44%) were male and 56 (56%) were female.
Table 1 shows background data for the 76 participants who were followed through to day 30. Induced abortion was performed on 62 viable fetuses of those participants, while 14 experienced spontaneous miscarriage. Eight fetuses of 14 were diagnosed by loss of fetal heart beat, and six were blighted ovum. Seventy participants were primigravidas, whereas the remaining six had previously given birth to a female child. Mean maternal age of the 76 participants was 24.3±6.2 years, and mean gestational age was 7.8±1.6 weeks. Mean age of spontaneous miscarriage women was significantly older than artificial abortion group (P<.05). No significant difference in mean gestational age was identified.
Table 2 presents information on sex of the chorion. The chorion was from a male fetus in 36 of 76 participants and from a female fetus in the remaining 40 participants. No significant differences were seen between these two participant groups in terms of maternal or gestational age.
Figure 1 shows time-course changes in the positive rate for microchimerism as a function of each fetal sex, judged on the basis of the presence or absence of SRY. On day 0, 19 of 36 participants (52.8%) who were pregnant with a male fetus were positive for microchimerism, but that number decreased to only 2 (5.6%) by day 7 and to 0 (0%) by day 30. Comparison of positive rates for microchimerism at those three time points of day 0, day 7, and day 30 showed a significant difference in the decrease (P<.001). In addition, none of the participants found to be negative for microchimerism on day 0 subsequently yielded positive test findings for microchimerism on day 7 or day 30. All participants from whom chorionic tissue indicated that the fetus had been female tested negative for SRY at each of the blood sampling times.
Fetal microchimerism occurs during pregnancy due to fetomaternal transfusion of blood cells across the placenta. The literature indicates that fetal cells are first detected in maternal blood after about gestational week 4,11 and the state of temporary microchimerism due to the presence of fetal cells in the maternal body continues until parturition. With regard to the time course for the disappearance of fetal cells from the maternal body after a normal birth, Hamada et al12 reported that fetal nucleated cells could no longer be detected in maternal peripheral blood at 3 months after parturition. In addition, Ariga et al11 reported that cell-free fetal DNA disappeared from the maternal circulation immediately after birth, whereas cellular fetal DNA disappeared by approximately 1 month postpartum. Conversely, no reports seem to have examined the disappearance of fetal cells from the maternal body after early termination of a pregnancy. The objective of the present study was thus to estimate fetal microchimerism after fetal loss in early pregnancy, the status of which has not yet been elucidated.
Our results revealed that, for women who had been confirmed to have been pregnant with a male fetus on the basis of chorionic testing, the detection rate for SRY-positive cells in peripheral blood before D&C of approximately one half decreased to 5.6% by day 7 after abortion and 0% by day 30. Moreover, none of the participants who were negative for SRY on day 0 later tested positive on day 7 or day 30. Unsurprisingly, all mothers who were confirmed to have been pregnant with a female fetus (serving as negative controls in this study) tested negative for SRY throughout the study period.
Comparison with the report of Ariga et al11 allows us to surmise that, in the case of women who had been pregnant with a male fetus, disappearance of fetal cells from the maternal body follows the same time course as seen after the birth of a male neonate. That is, fetal cells that have become mixed in the maternal body as a result of pregnancy or termination of pregnancy are eventually eliminated from the maternal blood (to be exact, levels decrease below the limit of detection). If this is true, new questions are raised: by what mechanism, and when, do fetal cells reappear in the maternal blood and establish “permanent microchimerism”? To date, we have no answers to these questions. In a 2004 report, Khosrotehrani et al13 suggested that fetomaternal microchimerism results from the establishment of fetal multilineage cells in the maternal body and acquisition of the capacity to differentiate.
Another paradox is that cells of male origin have been detected even in a certain percentage of women who have never given birth to a male child. Yan et al5 reported that the incidence of detecting DNA of male origin in women with no history of giving birth to a male child was 8% in women who had only given birth to a female child, 22% in women who had experienced spontaneous miscarriage, 57% in women who had undergone induced abortion, and 10% in women who had never been pregnant. Interestingly, the incidence of microchimerism is extremely high in women with a history of abortion. In addition, Yan et al reported that the blood of female infants and umbilical cord blood of female newborns were negative for microchimerism.
Imamura et al3 investigated the incidence of permanent microchimerism in nonpregnant women on the basis of the presence or absence of a history of pregnancy. They reported that the incidence of microchimerism in women who had given birth to a male child was 52%, whereas the incidence was 25% in women who had only given birth to a female child and 17% in nonpregnant women. In comparison, the present study found that 19 of 36 women (52.8%) who had been pregnant with a male fetus were positive for microchimerism on day 0. That incidence is almost the same as the incidence of 52% reported by Imamura et al3 for permanent microchimerism in women with a history of having given birth to a male child. Similarly, we detected microchimerism in 19 of 76 participants (25%) on day 0, and that incidence is also close to the incidences of 25% and 17% reported by Imamura et al3 for permanent microchimerism in women who, respectively, had only given birth to a female child or were not pregnant. The participant group of women for Imamura et al3 who had never given birth to a male neonate included women who had undergone induced abortion or experienced spontaneous miscarriage and also women for whom the historical status of miscarriage was unclear. For that reason, we can surmise that the incidence of microchimerism was increased in women who had never given birth to a male child. That is, the incidence of temporary microchimerism detected before D&C may reflect the incidence of permanent microchimerism reported by Imamura et al.3
One noteworthy finding of the present study was the fact that all women who had been pregnant with a female fetus were negative for microchimerism throughout the test period. In the literature,3,5 the positive rate for microchimerism was 10–17%, even in women who had never been pregnant. Moreover, Yan et al5 reported that the blood of female infants and umbilical cord blood of female newborns were negative for microchimerism. When that finding is taken together with our finding that all women who had carried a female fetus were negative for microchimerism, we can surmise that there is strong possibility that women with a male fetus who experience miscarriage or receive a chemical abortion will later become positive for microchimerism.
Wataganara et al14 reported that a comparison of surgical and medical termination at first trimester showed that the amount of fetal DNA in the maternal blood was higher in participants who had undergone surgery. This may be because surgical termination is invasive and thus causes destruction of trophoblastic villi or increases the extent of fetomaternal transfusion.
Given the present results, fetal cells that have once been eliminated from the maternal circulation after D&C, subsequently become detectable in the maternal circulation. In the case of miscarriage early in pregnancy, and particularly in the case of a history of surgical abortion, the positive rate of detection of microchimerism seems quite likely to be increased.
Both the duration after which fetal cells reappear in the maternal blood and the reason for this phenomenon remain unclear. Long-term follow-up of women who had been pregnant with a male fetus in an attempt to clarify if and when they become positive for SRY, whether even participants who were negative for microchimerism before surgical abortion also become positive at a later date, and whether any associations exist with later onset of some disease.
1. Lo YM, Lo ES, Watson N, Noakes L, Sargent IL, Thilaganathan B, et al. Two-way cell traffic between mother and fetus: biologic and clinical implications. Blood 1996;88:4390–5.
2. Bianchi DW, Zickwolf GK, Weil GJ, Sylvester S, DeMaria MA. Male fetal progenitor cells persist in maternal blood for as long as 27 years postpartum. Proc Natl Acad Sci U S A 1996;93:705–8.
3. Imamura S, Sato A, Ohto H. Pregnancy-induced microchimerism [in Japanese]. Fukushima J Med Sci 2001;51:113–9.
4. Lambert NC, Lo YM, Erickson TD, Tylee TS, Guthrie KA, Furst DE, et al. Male microchimerism in healthy women and women with scleroderma: cells or circulating DNA? A quantitative answer. Blood 2002;100:2845–51.
5. Yan Z, Lambert NC, Guthrie KA, Porter AJ, Loubiere LS, Madeleine MM, et al. Male microchimerism in women without sons: quantitative assessment and correlation with pregnancy history. Am J Med 2005;118:899–906.
6. Nelson JL, Furst DE, Maloney S, Gooley T, Evans PC, Smith A, et al. Microchimerism and HLA-compatible relationships of pregnancy in scleroderma. Lancet 1998;351:559–62.
7. Artlett CM, Smith JB, Jimenez SA. Identification of fetal DNA and cells in skin lesions from women with systemic sclerosis. N Engl J Med 1998;338:1186–91.
8. Evans PC, Lambert N, Maloney S, Furst DE, Moore JM, Nelson JL. Long-term fetal microchimerism in peripheral blood mononuclear cell subsets in healthy women and women with scleroderma. Blood 1999;93:2033–7.
9. McGraph H Jr. Elective pregnancy termination and microchimerism: comment on the article by Khosrotehrani et al. Arthritis Rheum 2004;50:3058–9.
10. Lo YM, Patel P, Sampietro M, Gillmer MD, Fleming KA, Wainscoat JS. Detection of single-copy fetal DNA sequence from maternal blood. Lancet 1990;335:1463–4.
11. Ariga H, Ohto H, Busch MP, Imamura S, Watoson R, Reed W, et al. Kinetics of fetal cellular and cell-free DNA in the maternal circulation during and after pregnancy: implications for noninvasive prenatal diagnosis. Transfusion 2001;41:1524–30.
12. Hamada H, Arinami T, Hamaguchi H, Kubo T. Fetal nucleated cells in maternal peripheral blood after delivery in cases of fetomaternal hemorrhage. Obstet Gynecol 1995;85:449–51.
13. Khosrotehrani K, Johnson KL, Cha DH, Salomon RN, Bianchi DW. Transfer of fetal cells with multilineage potential to maternal tissue. JAMA 2004;292:75–80.
14. Wataganara T, Chen AY, LeShane ES, Sullivan LM, Borgatta L, Bianchi DW, Johnson KL. Cell-free DNA levels in maternal plasma after elective first-trimester termination of pregnancy. Fertil Steril 2004;81:638–44.