Abstract
BACKGROUND: The objective of the study was to investigate the developmental fate of oocytes with increased cytoplasmic viscosity as assessed by the persistence of the injection funnel after withdrawal of the ICSI pipette. METHODS: For this purpose, 1008 oocytes showing a characteristic injection funnel during ICSI were subdivided into two groups according to the oocyte’s ability to restore its spherical shape within 2–3 min after ICSI. Fertilization and further development was evaluated in both groups. In addition, implantation and pregnancy rates were analysed. RESULTS: In the funnel positive cohort (group 1) significantly fewer oocytes degenerated after injection (P < 0.01) compared with oocytes without persistent funnel (group 2). However, at zygote stage, presence of a halo (P < 0.05) and a optimal pronuclear pattern 0 (P < 0.01) was increased in group 2. In addition, significantly fewer poor quality embryos were found in this group (P < 0.01). The number of good quality blastocysts but not blastocyst formation was increased in group 2 (P < 0.05). This resulted in an increased clinical pregnancy rate if embryos which derived exclusively from funnel negative oocytes were transferred (P < 0.05). CONCLUSIONS: Our data suggest that cytoplasm of higher viscosity delays development up to cleavage stage and impairs optimal development. Injection funnel persistence was found to be a negative prognostic marker of preimplantation development.
Introduction
Assisted reproductive technologies routinely use controlled ovarian hyperstimulation (COH) for oocyte recruitment and ovulation induction. Thus, an increased number of oocytes can be collected, though at the possible risk that not all gametes will show the same developmental competence. This heterogeneity is probably due to intrinsic differences in oocytes recovered either from follicles with sufficient blood supply or from follicles with diminished vascularization and subsequent hypoxia (Van Blerkom et al., 1997). Such oocytes may be characterized by an impaired synchronization of both nuclear and cytoplasmic maturation. Data from in‐vitro matured oocytes support the hypothesis that cytoplasmic maturation may be dissociated from nuclear maturation, indicating that though resumption of meiosis can be achieved, maturation of cytoplasm may be impaired (Mikkelsen and Lindenberg, 2001; Combelles et al., 2002).
Cytoplasmic maturation involves numerous metabolic and structural modifications in preparation for subsequent fertilization (Eppig et al., 1994). Since enucleated oocytes show similar maturation‐associated changes in protein synthesis as their nucleated counterparts (Sun and Moor, 1991) it is likely that cytoplasmic maturation is, at least in part, under cytoplasmic control (Eppig, 1996).
Deficient cytoplasmic maturity may be reflected by certain cytoplasmic abnormalities (e.g. cytoplasmic inclusions, vacuoles, smooth endoplasmic reticulum clustering) visible at light microscope level. In contrast with conventional IVF (Bedford and Kim, 1993) cytoplasmic dysmorphism is unlikely to have any impact on fertilization rate and embryo quality in ICSI patients (De Sutter et al., 1996; Balaban et al., 1998; Ebner et al., 2000) though ICSI of dysmorphic gametes can result in embryos with reduced developmental capacity (Alikani et al., 1995; Serhal et al., 1997).
In addition to cytoplasmic inclusions, apparent differences in cytoplasmic texture and density have been described (Kahraman et al., 2000; Meriano et al., 2001). Extensive cytoplasmic granularity may either be homogeneous, affecting the whole gamete, or centrally located. The latter was found to be negatively correlated with ongoing pregnancy rate (Kahraman et al., 2000). In contrast, slight or moderate granularity has been accepted as a normal feature of oocytes. It is conceivable that an increased viscosity of the cytoplasm may constrain cell organelles and/or pronuclei in their movement preventing the zygote from achieving alignment of both pronuclei or alignment of pronuclei with respect to the polar bodies, thereby severely impairing polarity and further preimplantation development (Edwards and Beard, 1997; Garello et al., 1999).
Though it has been reported that granular areas are more viscous than the surrounding cytoplasm (Payne et al., 1997) there is a lack of markers of increased cytoplasmic viscosity. However, everyday laboratory work reveals that not all oocytes are able to restore their spherical shape immediately after injection of a spermatozoon. This phenomenon is possibly due to a decreased fluidity of the cytoplasm, which may result in a reduced intracellular pressure. Therefore, this prospective study was set up to investigate the developmental fate of oocytes with impaired fluidity of the cytoplasm as assessed by the persistence of the injection funnel after withdrawal of the ICSI pipette.
Materials and methods
All 137 couples involved in this prospective study were referred to our IVF unit because of male subfertility. In 26 cases an additional tubal factor could be diagnosed. The mean age of all women was 32.2 ± 4.8 years (range 21–40 years). During the 2‐month period of evaluation all ICSI cycles were included except those performed on a weekend in which an accurate schedule of morphological assessment could not be ensured. Patients in whom the majority of oocytes exhibited sudden or difficult oolemma penetration during ICSI (Ebner et al., 2002) were excluded because either no injection funnel could be observed or the intensity of manipulation during the injection might have contributed to funnel persistence.
In preparation for oocyte collection COH was conducted using either a long protocol (n = 39) or an antagonist protocol (n = 98). In the long protocol down‐regulation of the pituitary was achieved with the GnRH agonist buserelin (Suprecur®; Aventis Pharma, Vienna, Austria). Stimulation was initiated with hMG (Menogon®; Ferring, Kiel, Germany).
In the GnRH antagonist protocol recombinant FSH (Puregon®; Organon, Vienna, Austria) was started on day 2 of the cycle. In addition, a GnRH antagonist (Orgalutran®; Organon) was administered after 5–6 days of stimulation, depending on the presence of a 12–13 mm follicle in the ultrasound scan.
In all patients ovulation was induced with 5000–10 000 IU hCG (Pregnyl®; Organon) provided that the lead follicle reached a diameter of 19 mm and serum estradiol appeared adequate. Oocyte retrieval was carried out transvaginally under ultrasound guidance 36 h after hCG administration.
After incubation for 2–4 h (BM1 medium; NMS Bio‐Medical, Switzerland) brief exposure to 80 IU/ml hyaluronidase (MediCult, Copenhagen, Denmark) facilitated mechanical removal of the cumulus cells. Following this procedure all ICSI procedures were carried out by the same embryologist according to our previously published guidelines (Ebner et al., 2001) with mechanical immobilization of the spermatozoa (×400 magnification) and the polar body being held at the 6 o’clock position. Documentation was carried out using imaging and archival software (Octax Eyeware®; MTG, Altdorf, Germany), thus minimizing inter‐individual differences in scoring.
Immediately prior to injection oocytes showing vacuoles or dark central granulation of the cytoplasm were separated from the others in order to minimize the possible influence of severe forms of cytoplasmic anomalies on ICSI outcome.
The remaining metaphase II (MII) oocytes showed no to moderate granulation with or without minor inclusions. Due to our selection criteria only MII oocytes showing normal response to the injection pipette with a resultant characteristic funnel prior to oolemma breakage were included in our study (Palermo et al., 1996; Ebner et al., 2002). Within 2–3 min after injection and withdrawal of the glass pipette mature oocytes were assessed for the presence of an injection funnel (Figure 1). To confirm persistence of the funnel the oocytes were rotated by means of the holding pipette to detect an invagination on the surface of the gametes. According to this behaviour oocytes were cultured in two different groups (BM1 medium).
Control for signs of an adequate fertilization process (two pronuclei, two polar bodies) was carried out 18–20 h post‐ICSI. Zygotes were checked for polarity (Garello et al., 1999), presence of a halo (Payne et al., 1997) and pronuclear pattern (Tesarik and Greco, 1999; Montag and Van der Ven, 2001). At this stage, the medium was changed to Blastassist System Medium 1 (MediCult).
Early embryo development (number of blastomeres and percentage of fragmentation) was evaluated 42–44 h after injection. From day 2 onwards, the first sequential medium was replaced by Blastassist System Medium 2 (MediCult) which was changed daily until day of transfer (day 3 or 5). Blastocyst quality on day 5 was assessed according to the degree of expansion and morphological criteria. In detail, good quality blastocysts were characterized by a tightly packed inner cell mass (ICM) and the formation of a cohesive trophectoderm (TE).
Transfer was either done on day 3 (n = 94) or on day 5 at the blastocyst stage (n = 42). In one patient, no blastocysts formed in culture and so transfer was cancelled. No priority was given to transferring embryos or blastocysts that derived from oocytes with less viscous cytoplasm. Selection in terms of transfer was exclusively based on morphological assessment. Consequently, approval of our Institutional Review Board was not sought.
All data of this prospective investigation were compared using χ2‐test, t‐test, and Mann–Whitney U‐test. Significance was defined as P < 0.05.
Results
A total of 1182 oocytes were collected from 137 patients. In 85.3% (1008/1182) of the gametes a first polar body was clearly visible. A total of 85 MII oocytes degenerated after injection (8.4%). The overall fertilization rate of 2PN (pronuclei) was 70.3% (709/1008).
Approximately one‐fifth (26/137) of all patients had no oocytes with a persistent funnel after withdrawal of the injection pipette, whereas nine (6.6%) exclusively showed such gametes. In a total of 355 oocytes the injection funnel did not vanish within 2–3 min (35.2% of all MII oocytes). There was no relation to stimulation protocol (P > 0.05) since frequency of a persistent funnel in the long protocol (36.9%) did not differ from that in the antagonist protocol (34.5%). Female age showed no correlation with funnel persistence.
Table I compares the outcome of the ICSI procedure on day 1 in funnel positive oocytes (group 1) with that in funnel negative ones (group 2). No differences in fertilization rate could be observed between oocytes with persistent injection funnel and oocytes without this feature. The latter group showed a significant increase in degeneration rate after ICSI when compared to their counterparts (P < 0.05).
Table II summarizes the morphological assessment at the zygote stage in both cohorts. Group 2 showed significantly more pronuclear patterns with good prognosis (pattern 0B) compared with group 1 (P < 0.01). In addition, gametes with immediate restoration of the spherical shape showed an increased incidence of halos (P < 0.05). There was no relationship between stimulation procedure and either presence of a halo or pattern 0 rate.
In Table III, morphological features of day 2 and developmental rate to day 5 have been compared. Overall fragmentation but not number of blastomeres was significantly increased in group 1 (P < 0.05). This result was probably due to a dramatic increase in the number of poor quality embryos compared with group 2 (P < 0.01). There was no relationship between cytoplasmic viscosity and blastocyst formation rate (P = 0.07). However, the number of good quality blastocysts was significantly higher in the cohort of oocytes without persistent funnel (P < 0.05).
Out of 136 transfers, 42 clinical pregnancies could be achieved (30.9%). Homogeneous transfers with respect to cytoplasmic fluidity could be compared in 87 cases (56 exclusively funnel negative, 31 funnel positive). Table IV summarizes the outcome of the ICSI procedure in case of homogeneous transfer. Clinical pregnancy rate was found to be significantly increased if embryos derived exclusively from funnel negative oocytes were transferred (P < 0.05). No difference in female age, abortion rate, multiple pregnancy rate or implantation rate could be found.
Discussion
Van Blerkom et al. (1997) have shown that at the time of ovulation induction fully grown follicles with comparable appearance can exhibit widely varying rates of vascularization. Resultant hypoxia in follicles with diminished or absent blood flow generated MII oocytes with developmental incompetence (Van Blerkom et al., 1997). While the actual consequences at cellular level still remain to be determined, the maturation process of the immature oocyte, e.g. synchronization of nuclear and ooplasmic maturation, is likely to be a critical event, being very sensitive to changes in follicular environment. Whereas resumption of meiosis can be assessed easily after extrusion of the first polar body, cytoplasmic maturation involving several processes that prepare the egg for activation, fertilization and preimplantation development (Eppig, 1996) may not be estimated at light microscopic level. Most of the cytoplasmic anomalies detectable with the help of a light microscope have been shown to be unrelated to preimplantation development in ICSI (De Sutter et al., 1996; Balaban et al., 1998; Ebner et al., 2000). However, to our knowledge, variations in cytoplasmic texture have not yet been followed through early development in vitro.
Intracellular pressure and ooplasm fluidity can be estimated by the extent to which the ooplasm rises within the injection pipette immediately following penetration of the oolemma prior to aspiration of cytoplasm. In addition, ooplasm of higher viscosity is more likely to adhere to the spike of the injection pipette which could be seen in a limited number (∼0.5%) of injections after withdrawal of the glass tool (Ebner et al., 2001). However, these observations are strictly empirical and cannot serve as prognostic markers. Persistence of the injection funnel after ICSI reflects a deficiency in cytoplasmic texture. Flux characteristics of cytoplasm are altered in the more viscous type which does not allow the oocyte to restore its original spherical shape as fast as seen in gametes with a more aqueous ooplasm.
The injection funnel is likely to be responsible for sealing the breach during injection (Kimura and Yanagimachi, 1995; Palermo et al., 1996). The absence of such a protective mechanism in oocytes with sudden breakage (without any injection funnel at all) causes an increase in oocyte degeneration (Palermo et al., 1996; Ebner et al., 2001). Since all oocytes involved in this prospective study showed an injection funnel during ICSI, the observed difference in degeneration rate (Table I) may be explained by a difference in cytoplasm fluidity. In other words, increased viscosity of the cytoplasm (persistent funnel) may keep it from leakage, whereas oocytes with more aqueous cytoplasm may tend to leak more frequently after ICSI.
Our data suggest that overall fertilization rate is not influenced by cytoplasmic texture in ICSI. However, since differences in ooplasm viscosity have been observed (Payne et al., 1997) it is possible that alterations in viscosity may delay abuttal of pronuclei by severely impairing microtubule organization. In contrast, a more fluid cytoplasm would promote optimal conditions in preparation for further preimplantation development.
Since there is some evidence that pronuclear alignment and alignment of nucleoli at the site of pronuclear contact are time‐dependent (Tesarik and Kopecny, 1989; Payne et al., 1997), this may have been manifested by a higher frequency of pronuclear patterns of good prognosis (Tesarik and Greco, 1999; Montag and Van der Ven, 2001) in group 2 (Table II). This finding is consistent with data from literature showing higher proportions of optimal pronuclear patterns in ICSI zygotes compared with IVF zygotes (Montag and Van der Ven, 2001) since direct placement of one single spermatozoon will shorten the fertilization process (Nagy et al., 1998). Our results with respect to pronuclear patterns suggest further subdivision of pattern 0 (Tesarik and Greco, 1999) as recommended by Montag and Van der Ven (2001).
Additional evidence supporting the hypothesis that increased viscosity hinders optimized cellular processes may be gained from the observation that significantly more zygotes showed a cytoplasmic flare in group 2 (Table II). This phenomenon is called cytoplasmic halo and is the manifestation of a microtubule‐mediated withdrawal of mitochondria and other cytoplasmic components to the perinuclear region (Scott and Smith, 1998). Oocytes with affected cytoplasm did not show this positive prognostic feature in terms of developmental competence as frequently as did oocytes with unaffected cytoplasm. It may be speculated that suboptimal cytoplasmic texture compromizes microtubule‐organized concentration of mitochondria to perinuclear regions and thus impairs cell cycle regulation severely.
According to the literature, our data suggest that optimal pronuclear patterns (0A, 0B) on day 1 are closely related to embryo morphology on day 2 (Balaban et al., 2001; Montag and Van der Ven, 2001; Salumets et al., 2001). Probably due to a dramatic increase in the rate of poor quality embryos (P < 0.01) the mean percentage of fragmentation was significantly higher in embryos from funnel positive oocytes (P < 0.05, Table III). One possible explanation for an increase in fragmentation might be that frequency and periodicity of cytoplasmic waves, which are known to be correlated to embryo quality (Payne et al., 1997), may have been affected by a higher cytoplasmic viscosity resulting in embryos of lower quality.
However, the similar blastocyst formation rates seen in both groups indicate that embryo morphology on day 2 or 3 may be an inappropriate prognostic factor in some patients (Rijnders and Jansen, 1998). Since approximately half of the embryos reached blastocyst stage in both cohorts it appears that embryos derived from oocytes with persistent funnel after ICSI may overcome the developmental delay found in earlier stages.
However, when considering exclusively those blastocysts which extended beyond the early blastocyst stage and showed a distinct ICM consisting of many cells and a cohesive trophectoderm, a significant difference between both groups could be observed (P < 0.05). In detail, two thirds were top quality blastocysts in the group 2 whereas <50% of the ‘affected’ embryos formed optimal blastocysts in vitro. This benefit is reflected by an significant increase in clinical pregnancy rate in patients who had homogeneous transfers of embryos or blastocysts that developed from oocytes without a persistent funnel compared with those who had the opposite concepti transferred.
To conclude, our data indicate that MII oocytes with an injection funnel persisting for some time after ICSI are likely to represent a cohort of gametes with an intrinsic ooplasmic defect having a negative influence on developmental competence. In our prospective study, we found numerous indications of an involvement of cytoplasmic viscosity in preimplantation development throughout the time of in‐vitro culture. However, this developmental disadvantage could be compensated for during subsequent cleavages as cleavage rate and blastocyst formation were not affected. Nevertheless, in the present study blastocyst quality as well as clinical pregnancy rates indicate that information regarding developmental competence is gained during the oocyte or zygote stage rather than at the cleavage stages.
Figure 1. (A) MII oocyte showing characteristic injection funnel after withdrawal of glass pipette. (B) Oocyte with immediate restoration of the spherical shape after ICSI. Arrows indicate sperm head and represent 10 µm.
Comparison of ICSI outcome between oocytes with persistent funnel and oocytes without
| Persistent funnel | No persistent funnel | P‐value | |
| No. of MII oocytes | 355 | 653 | |
| Fertilization (% 2PN) | 255 (71.8) | 454 (69.5) | NS |
| No fertilization (%) | 77 (21.7) | 116 (17.7) | NS |
| 1PN (%) | 3 (0.9) | 2 (0.3) | NS |
| 3PN (%) | 3 (0.9) | 14 (2.1) | NS |
| Degenerated (%) | 17 (4.8) | 68 (10.4) | <0.01 |
| Persistent funnel | No persistent funnel | P‐value | |
| No. of MII oocytes | 355 | 653 | |
| Fertilization (% 2PN) | 255 (71.8) | 454 (69.5) | NS |
| No fertilization (%) | 77 (21.7) | 116 (17.7) | NS |
| 1PN (%) | 3 (0.9) | 2 (0.3) | NS |
| 3PN (%) | 3 (0.9) | 14 (2.1) | NS |
| Degenerated (%) | 17 (4.8) | 68 (10.4) | <0.01 |
PN = pronucleus.
Comparison of ICSI outcome between oocytes with persistent funnel and oocytes without
| Persistent funnel | No persistent funnel | P‐value | |
| No. of MII oocytes | 355 | 653 | |
| Fertilization (% 2PN) | 255 (71.8) | 454 (69.5) | NS |
| No fertilization (%) | 77 (21.7) | 116 (17.7) | NS |
| 1PN (%) | 3 (0.9) | 2 (0.3) | NS |
| 3PN (%) | 3 (0.9) | 14 (2.1) | NS |
| Degenerated (%) | 17 (4.8) | 68 (10.4) | <0.01 |
| Persistent funnel | No persistent funnel | P‐value | |
| No. of MII oocytes | 355 | 653 | |
| Fertilization (% 2PN) | 255 (71.8) | 454 (69.5) | NS |
| No fertilization (%) | 77 (21.7) | 116 (17.7) | NS |
| 1PN (%) | 3 (0.9) | 2 (0.3) | NS |
| 3PN (%) | 3 (0.9) | 14 (2.1) | NS |
| Degenerated (%) | 17 (4.8) | 68 (10.4) | <0.01 |
PN = pronucleus.
Comparison of zygote morphology with respect to the persistence of the injection funnel
| Persistent funnel | No persistent funnel | P‐value | |
| No. of zygotes | 255 | 454 | |
| Pattern 0A (%) | 7 (2.7) | 26 (5.7) | 0.07 |
| Pattern 0B (%) | 51 (20.0) | 140 (30.8) | <0.01 |
| Pattern 1–5 (%) | 197 (77.3) | 288 (63.5) | <0.01 |
| Presence of halo (%) | 214 (83.9) | 407 (89.7) | <0.05 |
| Persistent funnel | No persistent funnel | P‐value | |
| No. of zygotes | 255 | 454 | |
| Pattern 0A (%) | 7 (2.7) | 26 (5.7) | 0.07 |
| Pattern 0B (%) | 51 (20.0) | 140 (30.8) | <0.01 |
| Pattern 1–5 (%) | 197 (77.3) | 288 (63.5) | <0.01 |
| Presence of halo (%) | 214 (83.9) | 407 (89.7) | <0.05 |
PMB = pronuclear membrane breakdown; PN = pronuclei.
Comparison of zygote morphology with respect to the persistence of the injection funnel
| Persistent funnel | No persistent funnel | P‐value | |
| No. of zygotes | 255 | 454 | |
| Pattern 0A (%) | 7 (2.7) | 26 (5.7) | 0.07 |
| Pattern 0B (%) | 51 (20.0) | 140 (30.8) | <0.01 |
| Pattern 1–5 (%) | 197 (77.3) | 288 (63.5) | <0.01 |
| Presence of halo (%) | 214 (83.9) | 407 (89.7) | <0.05 |
| Persistent funnel | No persistent funnel | P‐value | |
| No. of zygotes | 255 | 454 | |
| Pattern 0A (%) | 7 (2.7) | 26 (5.7) | 0.07 |
| Pattern 0B (%) | 51 (20.0) | 140 (30.8) | <0.01 |
| Pattern 1–5 (%) | 197 (77.3) | 288 (63.5) | <0.01 |
| Presence of halo (%) | 214 (83.9) | 407 (89.7) | <0.05 |
PMB = pronuclear membrane breakdown; PN = pronuclei.
Cleavage characteristics and blastocyst formation in oocytes with or without persistent injection funnel
| Persistent funnel (Group 1) | No persistent funnel (Group 2) | P‐value | |
| Day 2 | |||
| No. of cleavages | 252 | 449 | |
| No. of blastomeres | 4.1 ± 1.2 | 4.2 ± 1.4 | NS |
| Fragmentation (%) | 22.3 ± 20.2 | 16.1 ± 15.4 | <0.05 |
| No. of embryos without fragments (%) | 66 (26.2) | 131 (29.2) | NS |
| No. of bad quality embryos (%) | 48 (19.1) | 19 (4.2) | <0.01 |
| Day 5 | |||
| Total blastocyst formation (%) | 70/165 (42.4) | 114/218 (52.3) | NS (0.06) |
| Early blastocysts | 20 | 32 | NS |
| Blastocysts | 34 | 48 | NS |
| Expanded blastocysts | 11 | 23 | NS |
| Hatching blastocysts | 5 | 11 | NS |
| Top quality blastocyst | 19/50 (48.0) | 53/82 (64.6) | <0.05 |
| Persistent funnel (Group 1) | No persistent funnel (Group 2) | P‐value | |
| Day 2 | |||
| No. of cleavages | 252 | 449 | |
| No. of blastomeres | 4.1 ± 1.2 | 4.2 ± 1.4 | NS |
| Fragmentation (%) | 22.3 ± 20.2 | 16.1 ± 15.4 | <0.05 |
| No. of embryos without fragments (%) | 66 (26.2) | 131 (29.2) | NS |
| No. of bad quality embryos (%) | 48 (19.1) | 19 (4.2) | <0.01 |
| Day 5 | |||
| Total blastocyst formation (%) | 70/165 (42.4) | 114/218 (52.3) | NS (0.06) |
| Early blastocysts | 20 | 32 | NS |
| Blastocysts | 34 | 48 | NS |
| Expanded blastocysts | 11 | 23 | NS |
| Hatching blastocysts | 5 | 11 | NS |
| Top quality blastocyst | 19/50 (48.0) | 53/82 (64.6) | <0.05 |
Poor quality embryos include embryos with unevenly cleaved blastomeres and embryos with at least 50% fragmentation. Top quality blastocysts show distinct inner cell mass and cohesive trophectoderm (no early blastocysts included).
Cleavage characteristics and blastocyst formation in oocytes with or without persistent injection funnel
| Persistent funnel (Group 1) | No persistent funnel (Group 2) | P‐value | |
| Day 2 | |||
| No. of cleavages | 252 | 449 | |
| No. of blastomeres | 4.1 ± 1.2 | 4.2 ± 1.4 | NS |
| Fragmentation (%) | 22.3 ± 20.2 | 16.1 ± 15.4 | <0.05 |
| No. of embryos without fragments (%) | 66 (26.2) | 131 (29.2) | NS |
| No. of bad quality embryos (%) | 48 (19.1) | 19 (4.2) | <0.01 |
| Day 5 | |||
| Total blastocyst formation (%) | 70/165 (42.4) | 114/218 (52.3) | NS (0.06) |
| Early blastocysts | 20 | 32 | NS |
| Blastocysts | 34 | 48 | NS |
| Expanded blastocysts | 11 | 23 | NS |
| Hatching blastocysts | 5 | 11 | NS |
| Top quality blastocyst | 19/50 (48.0) | 53/82 (64.6) | <0.05 |
| Persistent funnel (Group 1) | No persistent funnel (Group 2) | P‐value | |
| Day 2 | |||
| No. of cleavages | 252 | 449 | |
| No. of blastomeres | 4.1 ± 1.2 | 4.2 ± 1.4 | NS |
| Fragmentation (%) | 22.3 ± 20.2 | 16.1 ± 15.4 | <0.05 |
| No. of embryos without fragments (%) | 66 (26.2) | 131 (29.2) | NS |
| No. of bad quality embryos (%) | 48 (19.1) | 19 (4.2) | <0.01 |
| Day 5 | |||
| Total blastocyst formation (%) | 70/165 (42.4) | 114/218 (52.3) | NS (0.06) |
| Early blastocysts | 20 | 32 | NS |
| Blastocysts | 34 | 48 | NS |
| Expanded blastocysts | 11 | 23 | NS |
| Hatching blastocysts | 5 | 11 | NS |
| Top quality blastocyst | 19/50 (48.0) | 53/82 (64.6) | <0.05 |
Poor quality embryos include embryos with unevenly cleaved blastomeres and embryos with at least 50% fragmentation. Top quality blastocysts show distinct inner cell mass and cohesive trophectoderm (no early blastocysts included).
Treatment outcome in homogeneous transfers with respect to the persistence of the injection funnel
| Funnel positive transfer | Funnel negative transfer | |
| Number of transfers | 31 | 56 |
| Positive βhCG (%) | 7/31 (22.6) | 23/56 (41.1) |
| Clinical pregnancies (%) | 5/31 (16.1)a | 20/56 (35.7)a |
| Multiple pregnancy rate (%) | 1/5 (20.0) | 3/20 (10.0) |
| Implantation rate (%) | 6/59 (10.2) | 23/107 (21.5) |
| Funnel positive transfer | Funnel negative transfer | |
| Number of transfers | 31 | 56 |
| Positive βhCG (%) | 7/31 (22.6) | 23/56 (41.1) |
| Clinical pregnancies (%) | 5/31 (16.1)a | 20/56 (35.7)a |
| Multiple pregnancy rate (%) | 1/5 (20.0) | 3/20 (10.0) |
| Implantation rate (%) | 6/59 (10.2) | 23/107 (21.5) |
aP < 0.05.
Treatment outcome in homogeneous transfers with respect to the persistence of the injection funnel
| Funnel positive transfer | Funnel negative transfer | |
| Number of transfers | 31 | 56 |
| Positive βhCG (%) | 7/31 (22.6) | 23/56 (41.1) |
| Clinical pregnancies (%) | 5/31 (16.1)a | 20/56 (35.7)a |
| Multiple pregnancy rate (%) | 1/5 (20.0) | 3/20 (10.0) |
| Implantation rate (%) | 6/59 (10.2) | 23/107 (21.5) |
| Funnel positive transfer | Funnel negative transfer | |
| Number of transfers | 31 | 56 |
| Positive βhCG (%) | 7/31 (22.6) | 23/56 (41.1) |
| Clinical pregnancies (%) | 5/31 (16.1)a | 20/56 (35.7)a |
| Multiple pregnancy rate (%) | 1/5 (20.0) | 3/20 (10.0) |
| Implantation rate (%) | 6/59 (10.2) | 23/107 (21.5) |
aP < 0.05.
References
Alikani, M., Palermo, G., Adler, A., Bertoli, M., Blake, M. and Cohen, J. (
Balaban, B., Urman, B., Sertac, A., Alatas, C., Aksoy, S. and Mercan, R. (
Balaban, B., Urman, B., Isiklar, A., Alatas, C., Aksoy, S., Mercan, R., Mumcu, A. and Nuhoglu, A. (
Bedford, J.M., and Kim, H.H. (
Combelles, C.M.H., Cekleniak, N.A., Racowsky, C. and Albertini, D.F. (
De Sutter, P., Dozortsev, D., Qian, C. and Dhont, M. (
Ebner, T., Yaman, C., Moser, M., Sommergruber, M., Feichtinger, O. and Tews, G. (
Ebner, T., Yaman, C., Moser, M., Sommergruber, M., Jesacher, K. and Tews, G. (
Ebner, T., Moser, M., Yaman, C., Sommergruber, M., Hartl, J., Jesacher, K. and Tews, G. (
Edwards, R.G. and Beard, H.K. (
Eppig, J.J., Schultz, R.M., O’Brian, M. and Chesnel, F. (
Eppig, J.J. (
Garello, C., Baker, H., Rai, J., Montgomery, S., Wilson, P., Kennedy, C.R. and Hartshorne, G.M. (
Kahraman, S., Yakin, K., Dönmez, E., Samli, H., Bahce, M., Cengiz, C., Sertyel, S., Samli, M. and Imirzalioglu, N. (
Kimura, Y. and Yanagimachi, R. (
Meriano, J.S., Alexis, J., Visram‐Zaver, S., Cruz, M. and Casper, R.F. (
Mikkelsen, A.L. and Lindenberg, S. (
Montag, M. and Van der Ven, H. (
Nagy, Z.P., Janssenswillen, C., Janssens, R., De Vos, A., Staessen, R., Van de Velde, H. and Van Steirteghem, A.C. (
Palermo, G., Alikani, M., Bertoli, M., Colombero, L.T., Moy, F., Cohen, J. and Z. Rosenwaks (
Payne, D., Flaherty, S.P., Barry, M.F. and Mathews, C.D. (
Rijnders, P.M. and Jansen, C.A.M. (
Salumets, A., Hydén‐Granskog, C., Suikkari, A.M., Tiitinen, A. and Tuuri, T. (
Serhal, P.F., Ranieri, D.M., Kinis, M., Marchant, S., Davies, M. and Khadum, I.M. (
Scott, L.A. and Smith, S. (
Sun, F.Z. and Moor, R.M. (
Tesarik, J. and Kopecny, V. (
Tesarik, J. and Greco, E. (
