Growth retardation in human blastocysts increases

Hum. Reprod. Advance Access published March 11, 2013 Human Reproduction, Vol.0, No.0 pp. 1– 8, 2013 doi:10.1093/humrep/det059

ORIGINAL ARTICLE Embryology

Growth retardation in human blastocysts increases the incidence of abnormal spindles and decreases implantation potential after vitrification IVF Namba Clinic, 1-17-28 Minamihorie, Nishi-ku, Osaka 550-0015, Japan *Correspondence address. Tel: +81-6-6534-8824; Fax: +81-66534-8876; E-mail: [email protected]

Submitted on December 13, 2012; resubmitted on January 19, 2013; accepted on February 13, 2013

study question: Does the human embryo growth rate affect the outcome of vitrified–warmed blastocyst transfer? summary answer: Following vitrification, the incidence of abnormal spindle morphology was increased and the implantation competence was decreased in growth-retarded embryos compared with normally developing embryos.

what is known already: Various types of spindle abnormality occur in human cleavage- and blastocyst-stage embryos. However, the incidence of abnormal spindle morphology in growth-retarded blastocysts is not known. Furthermore, there is conflicting data about the implantation potential of such blastocysts.

study design, size, duration: This was a retrospective cohort study including 878 single vitrified –warmed blastocyst transfers between 9 January 2010 and 10 July 2012, and an experimental study using 121 vitrified –warmed blastocysts donated to research. A comparison on the implantation potential and spindle shape of vitrified –warmed blastocysts was made between normally developing and growthretarded blastocysts.

participants/materials, setting, methods: In the clinical study, we compared the implantation rates of vitrified – warmed embryos that developed to the blastocyst stage on Day 5 after insemination (normally developing embryos) with those that required culture to Day 6 (growth-retarded embryo). In the experimental study, donated vitrified –warmed blastocysts were immunostained with an anti-a-tubulin antibody to visualize microtubules, an anti-g-tubulin antibody to image centrosomes and Hoechst 33342 or 4,6-diamidino-2phenylindole to visualize DNA. Confocal image analysis captured a z-series stack of 0.5-mm-thick optical sections encompassing the entire blastocyst. Only spindles with fusiform poles and with chromosomes aligned at the equator were classified as normal. main results and the role of chance: The implantation rate of growth-retarded embryos (47%, n ¼ 270) was significantly lower (P , 0.05) than that of normally developing embryos (57%, n ¼ 608). A total of 533 spindles were analyzed in Day 5 and 6 vitrified – warmed blastocysts. The incidence of abnormal spindles in the growth-retarded embryos (47%, n ¼ 274) was significantly higher (P , 0.01) than in the normally developing embryos (30%, n ¼ 259).

limitations, reasons for caution: Further studies are required to clarify the link between an increase in abnormal spindle formation and a decrease in embryonic implantation potential. wider implications of the findings: This study provided new insights into the possible implications of abnormalities in spindle formation in growth-retarded human blastocysts.

study funding/competing interest(s): Part of this work was supported by a grant from the Japan Society for the Promotion of Science (JPS-RFTF 23 580 397 to S.H.). No other competing interests are declared. Key words: blastocyst / growth retardation / embryo transfer / implantation / spindle formation

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Shu Hashimoto*, Ami Amo, Satoko Hama, Keijiro Ito, Yoshiharu Nakaoka, and Yoshiharu Morimoto

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Introduction

Materials and Methods Ethical approval This study was approved by the local ethics Institutional Review Board of IVF Namba Clinic. Vitrified embryos donated by couples who had completed their fertility treatment and gave informed consent were used for the in vitro study. For vitrified – warmed embryo transfer (ET), the couples received full explanations regarding the treatment and gave their consent to get involved.

Vitrification Embryos were cultured in the same sequential media all through the study period (Cleavage medium and Blastocyst medium; COOK Medical, Queensland, Australia) at 378C under 5% CO2, 5% O2 and 90% N2.

Embryos that developed to the blastocyst stage were equilibrated in 7.5% (v/v) ethylene glycol (EG, 054-0983, Wako Pure Chemical Industries, Ltd, Osaka, Japan), 7.5% (v/v) dimethyl sulfoxide (DMSO, D2650, Sigma-Aldrich, St. Louis, MO, USA), 20% (v/v) serum substitute supplement (SSS; 99193, Irvine Scientific, Santa Ana, CA, USA) and TCM 199 medium (12350-039, Invitrogen, Tokyo, Japan) in a range of 8 – 10 min, confirming shrinkage and re-expansion. They were transferred into vitrification solution consisting 15% (v/v) EG, 15% (v/v) DMSO, 1.0 M sucrose (192-00012, Wako Pure Chemical Industries, Ltd), 20% (v/v) SSS and TCM 199 medium. Each embryo was picked up with a small amount of vitrification solution (50 nl) and placed on top of a very fine polypropylene strip of a Cryotopw (Kitazato Corporation, Tokyo, Japan) and then immediately submerged into liquid nitrogen. Alternatively it was pipetted into the hole of a Rapid-iw (Vitrolife Sweden AB, Go¨teborg, Sweden) and introduced immediately into super-cooled air inside a straw held in liquid nitrogen and then sealed using an ultrasonic sealer as described previously (Hashimoto et al., 2013). These vitrification procedures were carried out within 90 s at room temperature. The survival rate after warming and the implantation potential for Rapid-i vitrification were the same as for Cryotop vitrification (Hashimoto et al., 2013). Thus, we considered Rapid-i vitrification in the same way as Cryotop vitrification.

Warming For warming, vitrified embryos were placed in 1 ml TCM 199 containing 20% SSS and 1 M sucrose, at 378C for 1 min, then diluted with 1 ml TCM 199 containing 20% SSS and 0.5 M sucrose and then diluted twice in 1 ml TCM 199 containing 20% SSS for 5 min each at room temperature. To obtain a numerical blastocyst morphology grading system based on that of Gardner and Lane (1997), the blastocyst grade before vitrification was converted to the compound blastocyst quality score (BQS, Rehman et al., 2007; Yamanaka et al., 2011). The BQS is a measure of blastocyst quality based on the established morphological criteria and is defined as the product of the degree of embryo development and of the inner cell mass and trophectoderm grades, where Grade A is given the value 3; Grade B is 2 and Grade C is 1. For example, a 3AB blastocyst has a BQS of 3 × 3 × 2, giving a total of 18.

Developmental competence after single ET To assess the developmental competence after ET of normally developing and growth-retarded blastocysts, 878 patients who underwent single vitrified – warmed blastocyst transfer under hormone-replacement cycles between 9 January 2010 and 10 July 2012 were included in the analysis (608 with normally developing and 270 with growth-retarded blastocysts); this did not include those receiving hatched blastocysts. Growth-retarded blastocysts were morula or early blastocyst stage: BL 1 (Gardner and Lane, 1997) on Day 5. Cryotop vitrification was carried out between 9 January 2010 and 10 July 2012 (n ¼ 828; ICSI cycles: 571, cIVF cycles: 257). Rapid-i vitrification was carried out between 14 January 2012 and 3 July 2012 (n ¼ 50; ICSI cycles: 35, cIVF cycles: 15). The proportion of normally developing blastocyst transfer cycles with Cryotop vitrification (69.0%, 571/828) was similar to that with Rapid-I vitrification (70%, 35/50). The proportion of ICSI cycles with normally developing blastocyst (67.6%, 411/608) was similar to that with growth-retarded blastocyst (72.2%, 195/270). There were 623 normally developing and 280 growth-retarded blastocysts warmed, and 608 (97.6%) and 270 (96.4%) were viable at 3 h post-warming, respectively. There was no difference in the survival rate between normally developing and growth-retarded blastocysts. The endometrium was prepared as described previously (Hashimoto et al., 2007) with some modifications by incremental doses


Normally, human embryos can reach the blastocyst stage of development on Day 5 after fertilization. However, there are often growth-retarded embryos that require 6 days to grow to this stage. Several studies using fresh blastocyst transfers have indicated that the rate of embryo development to the blastocyst stage affects the pregnancy outcome of in vitro fertilization (IVF) treatment cycles (Khorram et al., 2000; Shapiro et al., 2001; Barrenetxea et al., 2005), partly because of an inconsistency between the stage of embryo development and the receptivity of the recipient’s endometrium (Murata et al., 2005). On the other hand, studies on frozen –thawed or vitrified –warmed blastocyst transfers have reported conflicting results on whether the rate of blastocyst formation before cryopreservation affects treatment outcome (Marek et al., 2001; Behr et al., 2002; Hiraoka et al., 2004; Murata et al., 2005; Liebermann and Tucker, 2006; Richter et al., 2006; Levens et al., 2008; Shapiro et al., 2008; El-Toukhy et al., 2011). Moreover, a recent systematic review and meta-analysis concluded that those growth-retarded embryos that develop to the blastocyst stage by Day 6 have the same implantation potential as their Day 5 counterparts if the morphology of Day 6 blastocyst is similar to that of Day 5 blastocyst (Sunkara et al., 2010). On the other hand, the transfer of blastocysts showing early cavitation has produced more live offspring in cattle (Hasler et al., 1995) and the transfer of frozen–thawed Day 7 expanded bovine blastocysts has produced higher pregnancy rates than similar embryos requiring 8 days (Niemann et al., 1982). In addition, the rate of embryo development has been proposed as a good criterion for evaluation of the quality of the embryo produced in vitro; thus, embryos with a faster rate of division are developmentally more competent than their slower counterparts (Lemmen et al., 2008; Wong et al., 2010; Meseguer et al., 2011; Hashimoto et al., 2012). In this study, we compared the characteristics of vitrified –warmed human embryos that reached the blastocyst stage by Day 5 (normally developing) with those of embryos that required 6 days (growthretarded). We analyzed the implantation rates of the embryos after transfer, and evaluated spindle and chromosome configurations of donated blastocysts in vitro using confocal laser scanning microscopy to assess the effects of delayed blastocyst formation on the cytoskeleton.

Hashimoto et al.

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Abnormal spindles in growth-retarded embryos

Spindle abnormalities in blastocysts All donated blastocysts were vitrified using Cryotop vitrification. After warming, blastocysts were cultured for 18 h in KSOMaa medium (Biggers et al., 2000) containing 5% SSS, then fixed and immunostained. Briefly, the blastocysts were fixed in Microtubule Stabilization Buffer Extraction Fixative (MTSB XF, Albertini et al., 1984) at 378C for 10 – 15 min. The MTSB XF consisted of 0.1 M PIPES, 50% (v/v) deuterium oxide, 0.01% (v/v) aprotinin, 1 mM Taxol and 0.5% (v/v) Triton-X 100. It contained 2% (w/v) formaldehyde to stabilize the blastocyst’s cytoskeleton. Fixed blastocysts were washed in Ca++ /Mg++ -free phosphate-buffered saline (PBS) containing 1% (w/v) bovine serum albumin (PBS – BSA; BSA, A7638, Sigma-Aldrich) and then cultured in PBS – BSA at 378C for 60 min. The blastocysts were then triple-stained to visualize spindle poles, microtubules and DNA. Samples were incubated in a solution of primary mouse monoclonal antibody specific for g-tubulin (1:1000; T6557, Sigma-Aldrich) for 1 h at 48C to identify the spindle poles. After being washed in BSA – PBS three times, they were incubated in a secondary antibody: Alexa Fluorw 594-labeled Goat Anti-Mouse IgG (H+L) (1:1000; A-11005, Life TechnologyTM , Carlsbad, CA, USA) for 1 h at 48C. After further washes in BSA – PBS three times, they were incubated in Alexa488w-conjugated mouse anti-a-tubulin (1:100; 322588, Life TechnologyTM ) for 1 h at 48C to identify the microtubules. After three washes in BSA – PBS, they were stained for DNA with 10 mg/ml of bisbenzimide H 33342 trihydrochloride (Hoechst 33342, 591-01721; Wako Pure Chemical Industries Ltd) or 1 mg/ml 4,6-diamidino-2-phenylindole (DAPI; 340-07971; Wako Pure Chemical Industries Ltd) and mounted on slides in Vectashield antifade medium (Vector Laboratories, Burlingame, CA, USA) under a coverslip. The coverslips were then sealed with nail varnish. The samples were examined with a laser scanning confocal microscope (CellVoyagerTM CV1000; Yokogawa Electronic, Tokyo, Japan).

Confocal image analysis was typically accomplished by capturing a z-series stack of 0.5-mm-thick optical sections encompassing the entire blastocyst (Fig. 1). The images were acquired sequentially using the 561-nm wavelength of a light-emitting diode (LED) to image Alexa 594, the 488 nm wavelength of a LED to image Alexa 488 and the 405 nm laser wavelength of a LED to visualize Hoechst or DAPI staining of DNA. A 40 × oil immersion lens was used for imaging.

Classification of spindle abnormalities All interphase nuclei and metaphase/anaphase spindle and chromosome configurations were examined carefully and counted in each blastocyst. The criteria for classifying spindle abnormalities were as described by Chatzimeletiou et al. (2005, 2012). A spindle with astral-shaped or fusiform poles and with chromosomes aligned at the equator was classified as normal (Fig. 1). A spindle with one or two poorly defined or apparently absent poles, generally with misaligned chromosomes, was classified as having an abnormal shape. Spindles with more than two clearly defined astral poles and the characteristic ‘Y’- or ‘X’-shaped arrangement of Hoechst- or DAPI-labeled chromosomes were classified as multi-polar.

Statistical analysis Differences between the two groups were compared using the Mann – Whitney non-parametric U-test. Data are represented as the mean + SEM. Statistical analysis was performed using StatView version 5 (SAS Institute Inc., Cary, NC, USA), and P , 0.05 was considered significant.

Results Implantation potential A total of 878 vitrified– warmed human blastocysts were transferred. Of these, 608 embryos had developed to the blastocyst stage on Day 5 after insemination (normally developing) and 270 embryos had developed to this stage on Day 6 (growth-retarded). There were no differences in the blatocyst quality score (normally developing group, 18.0 + 0.3 versus growth-retarded, 18.6 + 0.6), in the mean female age (normally developing group, 34.7 + 0.1 years versus growth-retarded, 34.8 + 0.2 years), in the mean recipient endometrial thickness (normally developing group, 11.5 + 0.1 mm versus growthretarded, 11.3 + 0.1 mm) or in the number of ETs in past treatment (normally developing group, 1.6 + 0.0 versus growth-retarded 1.7 + 0.1) between Day 5 and 6 blastocyst transfers. The implantation rate (57.2 + 2.0%) and the ongoing pregnancy rate over 10 weeks (46.4 + 2.0%) from transfers of the normally developing blastocyst were both significantly higher (P , 0.05) than that of the growth-retarded blastocyst transfers (implantation rate 47.0 + 3.0% and ongoing pregnancy rate 38.9 + 3.0%; Fig. 2). The incidence of chromosomal abnormalities of abortuses from normally developing blastocyst transfers was 75% (24/32), similar to that of abortuses from growth-retarded blastocyst transfers (71%; 10/14).

Births As shown in Table I, there were no significant differences between normally developing (Day 5) and growth-retarded (Day 6) blastocysts in the proportion of male babies (normally developing 50.9% versus growth-retarded 52.9%), in the birthweight of babies (normally developing 3141 g versus growth-retarded 3046 g), in the gestational age


of oral estradiol valerate (Progynovaw; Bayer Schering Pharma Co., Ltd, Zu¨rich, Switzerland) from 1 to 4 mg for 2 weeks following administrations of GnRH agonist (600 mg/day, Suprecurw nasal solution 0.15%; Mochida Pharmaceutical Co., Ltd, Tokyo, Japan) for 3 weeks. Chlormadinone acetate (6 mg/day Lutoralw, Shionogi & Co., Ltd, Osaka, Japan) was administered after confirming the endometrial thickness to be more than 8 mm by ultrasonography. The daily doses of 3 mg estradiol valerate and 6 mg chlormadinone acetate were maintained until the pregnancy test. For women with positive pregnancy tests, transcutaneous estradiol patches (2.88 mg every 2 days, Estranaw, Hisamitsu, Saga, Japan) and transvaginal progesterone (400 mg of progesterone/day, Utrogestanw 200 mg, Ferring Pharmaceuticals Ltd, West Drayton, UK) were administered until 9 weeks of gestation. Progesterone (Progeston depotw 125 mg, Fuji Pharma Co., Ltd, Toyama, Japan) was administered intramuscularly on the day of ET with two additional injections after confirming pregnancy. Blastocyst transfer was carried out on the fifth day of Lutoral administration. Implantation was determined from the detection of a single intrauterine gestational sac by transvaginal ultrasound around 3 weeks after ET. Ongoing pregnancy was defined as a pregnancy developing beyond 10 weeks of gestation with a fetal heart beat confirmed using ultrasonography. The birthweight, the gestational age and the incidence of any peromelus were assessed in babies born. In this study, monozygotic twins were observed in two cycles (0.6%) of normally growing blastocyst transfer and in two cycles (1.5%) of growth-retarded blastocyst transfer, respectively. In the case of any abortion, the karyotype of the abortus was analyzed as described (Hashimoto et al., 2008). Slide preparations and G-banding of chromosomes were conducted according to standard protocols (Rooney and Czepulkowski, 1992).

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Figure 1 Maximum intensity projections of confocal sections (A – C) and confocal sections captured every 0.5 mm along the z-axis (D – L) of vitrified human blastocysts showing a normal astral bipolar spindle (long arrow), (C) a monopolar spindle with three g-tubulin spots (short arrow), an abnormally shaped spindle with four g-tubulin spots (arrowheads) and a multipolar spindle (*). Staining: DNA (blue), a-tubulin (green) or g-tubulin (red). (A) DNA, a-tubulin and g-tubulin. (B) a-tubulin and g-tubulin. (C) DNA and g-tubulin.

(normally developing, 275.7 days versus growth-retarded, 275.0 days) or in the incidence of congenital limb deformations (normally developing, 3.0%, n ¼ 231 versus growth-retarded, 2.4%, n ¼ 85).

Spindle abnormalities in blastocysts Vitrified –warmed human blastocysts were processed for cytoskeletal analysis of normally developing (n ¼ 67) and growth-retarded blastocysts (n ¼ 54). As shown in Table II, there was no difference in the mean female age between the normally developing (33.5 + 0.4 years) and growth-retarded blastocysts (33.8 + 0.4 years).

Spindles were observed in 64 of the normally developing blastocysts (95.5%) and 53 of the growth-retarded blastocysts (98.1%). The range of mitotic spindles was 0 –15 per blastocyst. The mean cell number of normally developing blastocysts (124.6 + 5.5) was significantly less than that of their growth-retarded counterparts (159.1 + 8.6, P , 0.01). Cytoskeletal analysis revealed that the proportion of blastocysts with only normal spindles on Day 5 (19/67; 28%) was significantly higher than that on Day 6 (7/54; 13%, P , 0.05). For the 64 normally developing blastocysts, 259 spindles were analyzed, of which 182 (70.3 + 2.8%) were normal, 28 (10.8 + 0.2%) were abnormally

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shaped, 28 (10.8 + 0.2%) had multipolar spindles and 21 (8.1 + 0.2%) had monopolar spindles (Table II and Fig. 3). For the 53 growth-retarded blastocysts, 274 spindles were analyzed, of which 144 (52.6 + 3.0%) were normal, 51 (18.6 + 2.4%) were abnormally shaped, 24 (8.8 + 1.7%) were multipolar and 55 (20.4 + 2.4%) were monopolar. The incidence of normal spindles in normally developing blastocysts after vitrification was significantly higher (P , 0.01) than in their growth-retarded counterparts. The incidences of monopolar and of abnormally shaped spindles in normally developing blastocysts were significantly lower (P , 0.01) than in their growth-retarded counterparts. Chromosomes arranged in a rosettelike pattern at the periphery of the monopolar microtubule array were often observed in the growth-retarded blastocysts (Fig. 3).

ab

P , 0.01 and cdP , 0.05 by Mann –Whitney non-parametric U-test.

These data clearly indicate that the incidence of abnormal spindles was increased and the implantation competence was decreased following vitrification in growth-retarded blastocysts compared with normally developing blastocysts. Detailed examination by confocal laser scanning microscopy revealed a significantly higher incidence of spindle abnormalities in the growth-retarded embryo group compared with normally developing embryos. The spindle aberrations included monopolar and abnormally shaped spindles. In particular, the incidence of monopolar spindles was greatly increased in growth-retarded embryos. Such spindles are frequently observed phenotypes that can result from mutations or from the inhibition or depletion of a large number of different proteins (Tillement et al., 2009). Normally, mitotic cells containing monopolar spindles show delayed mitosis caused by checkpoint control mechanisms until the spindle assembly defects are corrected (Paoletti et al., 1997). If correct assembly cannot be achieved, cells might die in mitosis or undergo checkpoint slippage. In this, they exit mitosis with eventual problems in chromosomal separation (Rieder and Palazzo, 1992; Paoletti et al., 1997; Musacchio and Hardwick, 2002) because checkpoint proteins at the kinetochores produce signals to delay chromosomal separation until all chromosomes are bioriented and aligned symmetrically between the two spindle poles (Ciliberto and Shah, 2009). The resulting daughter cells are often arrested in the next cell cycle or die from apoptosis (Paoletti et al., 1997). Thus, it is likely that an increase in monopolar spindles in embryos will lead to a decrease in implantation potential because of a drop in overall cell numbers through mitotic arrest. The proportion of chromosomal aberrations in abortuses in the case of growth-retarded blastocyst transfer was at a similar level as for abortuses from the transfer of normally developing blastocysts. In other words, the increase in spindle abnormalities in growthretarded blastocysts decreased their implantation potential, but did not affect the incidence of chromosomal aberrations in abortuses. Moreover, there were no significant differences between normally

Table I Perinatal data of normally developing and growth-retarded embryos. Birthweight

Proportion of male babies

Gestational age

Incidence of peromelus

............................................................................................................................................................................................. Normally developing blastocyst

3141 + 31 g (227)

50.9 + 3.3% (230)

275.7 + 0.8 days (227)

3.0 + 1.1% (231)

Growth-retarded blastocyst

3046 + 58 g (84)

52.9 + 5.4% (87)

275.0 + 1.6 days (83)

2.4 + 1.7% (85)

Table II Spindle abnormalities in normally developing and growth-retarded embryos. Female donor age

Number of cells

Proportions of

........................................................................................................ Normal spindle

Abnormal shape spindle

Multipolar spindle

Monopolar spindle

............................................................................................................................................................................................. Normally developing blastocyst

33.5 + 0.4

124.6 + 5.5a

70.3 + 2.8a

10.8 + 1.9a

10.8 + 1.9

8.1 + 1.7a

Growth-retarded blastocyst

33.8 + 0.4

159.1 + 8.6b

52.6 + 3.0b

18.6 + 2.4b

8.8 + 1.7

20.4 + 2.4b

ab

P , 0.01 by Mann –Whitney’s U-test.


Figure 2 Implantation potential of growth-retarded blastocysts.

Discussion

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Figure 3 Maximum intensity projections of confocal sections of an abnormal spindle (A – C) and three different monopolar spindles (D – F, G– I and J– L). Staining: Hoechst 33342 (blue), a-tubulin (green) or g-tubulin (red). (A, D, G and J) DNA, a-tubulin and g-tubulin. (B and K) DNA and a-tubulin. (C, F, I and L) DNA and g-tubulin. (E and H) a-tubulin and g-tubulin.

developing and growth-retarded blastocysts in the birthweight of babies, the gestational age or the incidence of peromelus. Taken together, it appears likely that most blastomeres with abnormal spindles are eliminated before implantation. A systematic meta-analysis based on the case of multiple embrto transfer cycles concluded that growth-retarded embryos, which developed to the blastocyst on Day 6, have the same implantation potential as normally developing counterparts if the morphology of Day 6 blastocyst is similar to that of Day 5 blastocyst (Sunkara et al., 2010). In the present study, the implantation potential after the transfer of single growth-retarded blastocysts was decreased despite there being no differences between normally developing and

growth-retarded blastocysts in the female’s background or in the BQS. The primary difference between previous reports and ours is the difference in the number of embryos transferred. Multiple blastocyst transfers may have led to analytical errors in previous studies. Our data also show that vitrification of growth-retarded blastocyst is practical for clinical application, consistent with several previous reports (Marek et al., 2001; Behr et al., 2002; Hiraoka et al., 2004; Murata et al., 2005; Liebermann and Tucker, 2006; Richter et al., 2006; Levens et al., 2008; Shapiro et al., 2008; El-Toukhy et al., 2011). However, the implantation potential was significantly reduced in growth-retarded blastocyst even though they showed almost the same morphology and quality as normally developing blastocysts.


There is no need to eliminate growth-retarded blastocysts from ET in clinical practice; however, we must remember that growth retardation indicates a decrease in embryo potential. Further investigation is warranted to elucidate the rate of development and the fate of embryos with abnormal spindles and how these might influence the chromosomal constitution of the developing blastocysts. We suggest that growth-retarded blastocysts can be used safely in clinical treatment, while considering any potential risks arising from abnormal divisions of affected spindles.

Authors’ roles S.H. was involved in the literature review, experimental design, data acquisition, interpretation and analysis, and manuscript preparation. A.A. was involved in the immunostaining procedure. S.H., K.I., Y.N. and Y.M. were involved in the clinical embryo vitrification and ET.

Part of this work was supported by a grant from the Japan Society for the Promotion of Science (JPS-RFTF 23580397 to S.H.).

Conflict of interest None declared.

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Funding

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