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Sensing cell membrane biophysical properties for detection of high quality human oocytes Zhongrong Chen, Zhiguo Zhang, Xiaojie Guo, Kashan Memon, Fazil Panhwar, Meng Wang, Yunxia Cao, and Gang Zhao ACS Sens., Just Accepted Manuscript • DOI: 10.1021/acssensors.8b01215 • Publication Date (Web): 25 Dec 2018 Downloaded from http://pubs.acs.org on December 26, 2018

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Sensing cell membrane biophysical properties for detection of high quality human oocytes Zhongrong Chen†, #, Zhiguo Zhang‡,

∇, #,

Xiaojie Guo§, #, Kashan Memon†, Fazil Panhwar†, Meng

Wang†, Yunxia Cao*, ∇, ‡, Gang Zhao*, † †Department

of Electronic Science and Technology, University of Science and Technology of China,

Hefei 230027, Anhui, China; ‡Reproductive

Medicine Center, Department of Obstetrics and Gynecology, the First Affiliated Hospital

of Anhui Medical University, Hefei 230022, Anhui, China; ∇Anhui

Province Key Laboratory of Reproductive Health and Genetics, Biopreservation and Artificial

Organs, Anhui Provincial Engineering Research Center, Anhui Medical University, Hefei 230022, Anhui, China; §Hefei

Blood Center, Hefei 230031, Anhui, China.

* Correspondence should be addressed to: Gang Zhao, Ph.D. Department of Electronic Science & Technology University of Science and Technology of China Road Jinzhai 96, Hefei 230027, Anhui, P. R. China Tel: +86-18256929838 E-mail: [email protected] (G.Z.) or [email protected] (Y.C.)

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ABSTRACT Oocyte quality plays a crucial role in the early development and implantation of the embryos, and consequently has a profound impact on the accomplishment of assisted reproductive technology (ART). Simple and efficient method for detecting high-quality human oocytes is urgently needed. However, the clinically used morphological method is time-consuming, subjective and inaccurate. To this end, we propose a practical and effective approach for detecting high-quality oocytes via on-chip measurement of the oocyte membrane permeability. We found that oocytes can be divided into two subpopulations (high-quality versus poor-quality oocytes) according to their membrane permeability differences, and as was further confirmed by subsequent in vitro fertilization (IVF) and development experiments (the blastocyst rates of high-quality and poor-quality oocytes were 60% and 0%, respectively). This approach shows great potentials in improving the success of ART, including both the fertilization and development rates, and thus it may have wide applications in the clinic.

KEYWORDS: Oocyte detection, Microfluidic device, High-quality human oocyte, Membrane permeability, Assisted Reproductive Technology (ART)

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With the development of reproductive medicine, assisted reproductive technology (ART) has gained growing attention from infertile couples and allowed the pregnancy of millions of infertility patients in the world.1 It is well known that the quality of human oocyte plays a crucial role in the initial embryo development and thus has a profound impact on the accomplishment of ART.2 High-quality human oocytes detection can not only increase the number of available embryos during ART treatment, but also optimize the transplantation process and promote the clinical pregnancy rate.3 Therefore, high-quality human oocyte identification is critical in the field of assisted reproductive medicine.4 In the clinic, the quality of human oocytes for fertilization is mainly determined using morphological characterization.5 The clinicians select the oocytes for fertilization by some particular indicators, such as the shape, the size, the first body integrity6 and the zona pellucida.7 However, this way is very subjective and inaccurate and can lead to misinterpretation of human oocyte quality.8 To solve this problem, image processing and polarized light (PolScope) technique were studied to improve the reliability and accuracy of the morphological method.9,

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Additionally, electron, confocal and

fluorescence microscopy combining with agents like dyes, fixative, etc. have also been tried to improve morphological analysis.11-13 Apart from that morphological method, there are some new investigations.14-17 Wherein, optical spectroscopy appears to evaluate the oocyte quality.18,

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Vidberg et al. presented a simple optical

micro-system for measuring the transmission spectra of oocytes to determine its maturation degree.18 Another prospective method focuses on the mechanical characteristics of oocyte, investigations proved that there is a link between the stiffness of oocytes and their biological properties.20,

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Thus, the

mechanical characterizations can be used for oocytes selection.22 In a recent study, Yenez et al. have found that there is a difference between the viscoelastic properties of viable and non-viable fertilized 3


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oocytes.23 They also demonstrated the mechanical stiffness of mouse embryo is a decisive criterion for detecting embryo quality.23 To select high-quality oocytes by the mechanical characteristics, diverse devices were developed. Liu et al. and Wacogne et al. developed a system that compresses the oocyte against several ﬂexible beams by a holding pipette to detect oocyte quality.24, 25 A nano-force sensor and a new force sensing platform using passive magnetic springs were presented to measure mechanical characteristics of oocytes.22, 26 It should be pointed out that these studies need complex equipment and are dependent on the experimental environment.22, 26 More recently, the genetic screening method was reported to optimize oocyte development rate and embryo quality.27-29 However, the cost of genetic screening is high, and the process is complicated.30 Currently, the permeability parameter measurement has become an effective way to assess the viability and function of cells or tissues,31, 32 especially using microfluidic techniques.33-39 This may be a novel way to detect high-quality oocytes. From the above, there is still a lack of a simple and efficient method for detecting high-quality oocyte. Here, we propose a simple and effective method of on-chip cell membrane permeability sensing for detecting high-quality human oocytes. We examined the permeability of inseminated-unfertilized oocyte membranes under different conditions using a self-made microfluidic device. Comparing with normal oocytes, we found that oocytes can be divided into two subpopulations based on the membrane permeability. Subsequent in vitro fertilization (IVF) and developmental experiments further confirmed that the two subpopulations correspond to high-quality and poor-quality oocytes, respectively. Given this, we develop a novel, simple and effective on-chip detection method to detect high-quality oocytes. It enables to markedly improve the fertilization and development rate of oocyte and promote the clinical pregnancy rate.

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 EXPERIMENTAL SECTION Reagents and ethical statement. All reagents were purchased from Sigma (St. Louis, MO, USA) unless otherwise specified. Gonadotropin-releasing hormone antagonist (GnRHa) was purchased from Ipsen Pharma (Boulogne-Billancourt, France). Recombinant human follicle-stimulating hormone (r-hFSH) and human chorionic gonadotropin (hCG) were purchased from Merck Serono Co. (Geneva, Switzerland). The isotonic solution (263 ± 0 mOsm) was comprised of Quinn's Advantage Medium with HEPES (In Vitro Fertilization, Inc., Trumbull, CT, USA) and 10% v/v serum substitute supplement (Irvine Scientific, Santa Ana, CA, USA); The hypertonic solutions: 1 M Ethylene Glycol (EG, 1440 ± 1 mOsm) or 1 M Propylene Glycol (PG, 1433 ± 1 mOsm) in the isotonic solution. 1 M CPA was chosen because it is typically used in cell slow freezing and permeability measurement. The study was approved by the Biomedical Ethics Committee of Anhui Medical University (Anhui, China). Source and preparation of oocytes. The mature human oocytes were obtained from patients who underwent IVF treatment at the First Affiliated Hospital of Anhui Medical University. Written consent was obtained from all subjects. The patients’ ages were less than 35 years old and received an ovarian stimulation (GnRHa and r-hFSH) to induce superovulation. When the size of 2-3 leading follicles on the ovary reached ≥18 mm in diameter as seen by ultrasound scan, 10,000 IU of hCG was given, and oocytes retrieval were performed 36 hours later. Oocytes were collected using single lumen 17G needle (Wallace, SIMS, UK) through the transvaginal ultrasound-guided route. The cumulus-oocyte complexes (COCs) in follicular fluid were isolated and picked up under a microscope (IX 71, Olympus, Japan) and then placed into the fertilization medium (COOK MEDICAL 5


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INC., Bloomington, USA) for 4 to 6 hours of culture in vitro at 37 °C with 6% CO2. Subsequently, the insemination was conducted by adding each patient’s husband sperm into the fertilization medium containing COCs. About 16~18h later, the oocytes were removed completely from cumulus and corona cells using fine bore glass pipettes and fertilization was assessed under an inverted microscope (IX 71, Olympus, Japan). The fertilized oocytes were placed into cleavage medium (COOK MEDICAL INC., Bloomington, USA) for further embryo culture, while the inseminated-unfertilized oocytes were collected for the membrane permeability study. For screening high/poor-quality oocytes, the oocytes isolated from COCs were collected to use directly. Design and fabrication of the microperfusion system. We designed a specialized microfluidic perfusion device for detecting high-quality oocytes. The whole system is shown in Figure 1A. It mainly includes microperfusion chamber chip, monitoring system, temperature control system and fluid control system. For monitoring system, the whole assembled structure is placed on a microscope (BX53, Olympus, Japan) and equipped with a CCD (DP 71, Olympus, Japan), they allied with a computer for recording the volume responses of the oocyte. A 150 × 50 × 10 mm rectangular chamber is mounted under the chip via silicone rubber. It is connected to a constant temperature water bath (NCB-2400, Eyela Tokyo Rikakikai Co., Ltd., Tokyo, Japan) for coolant flow and as a temperature-controlled stage. Among the microchannel, there is a narrow groove for locating the thermocouple (TT-K-36, OMEGA Engineering, INC, Stamford, USA). The rectangular chamber, water bath, and thermocouple composed the temperature control system. The fluid control system is comprised of three high-precision programmable syringe pumps (WK-101P, Nanjing Anerke Electronics Technology Co. Ltd., China), of which two are used to inject the perfusion solution and the other is used for collecting waste solution. The injection program of the isotonic solution is: 12 μl/min for 4 min and then paused for 12 min. For 6


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hypertonic solution, its injection program is the opposite of the isotonic solution. The operating speed of the pumps is 12 μl/min, such slow mixing method was selected to avoid the oocyte deformation. A polytetrafluoroethylene (PTFE) inlet tube is linked to two pumps via a “T” type three-way tube. Moreover, all pipelines of the microperfusion system are made from PTFE tube. Its thick and rigid wall can ensure the fluid flow is stable throughout the entire system.

Figure 1. Schematic illustration of the microfluidic perfusion system. (A) The composition of the whole system. (B) The 2D view of the microfluidic perfusion device and experimental principle. (C) The 3D-disassembled view of the microfluidic perfusion device. Note: drawing not to scale.

As shown in Figure 1B, the microperfusion chamber is a sandwich unit, two pieces of silica gel thin film (50-μm- and 100-μm-thick) with a microchannel are sandwiched between two pieces of silica glass. The silica gel thin films stick together tightly and do not permit leakages of solution. The microchannel is a two-layer structure, the lower level is made of two discontinuous rectangular microchannels (~50 7


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mm long and ~2 mm wide) cut from the 100-μm silica gel thin film, and the upper layer microchannel is ~110-mm long and ~2-mm wide cut from the 50-μm silica gel thin film. Two small holes of ~5-mm in diameters are drilled in the upper glass as inlet and outlet of the solution. A 3D-disassembled view of microperfusion chip is presented in Figure 1C. There is a barrier in the lower film and the upper channel is straight, the barrier is cut into an arc shape to capture the oocyte at the center of the channel. The channel in the upper layer is aligned with the lower channel, and all the components are clamped tightly with the file clips. These features make the microperfusion chamber particularly suitable for measuring the oocyte. Oocyte perfusion experiment and image analysis. Prior to each experiment, it was necessary to eliminate air bubbles and ensure that all pipelines were unobstructed. The temperature in the chamber was set at a predetermined value in advance using the constant temperature water bath. Then, the oocyte was injected smoothly into the chamber using a 1,000-μl pipette tip, it could be watched as it flowed through the channel. The injection speed of the syringe pump was 1 μl/min to avoid flushing out the oocyte. Once the oocyte was trapped, the perfusion solutions were switched with the syringe pumps to an isotonic solution for 4 min followed by a hypertonic solution for 12 min. Four typical temperatures (25, 15, 4 and 0 °C) were chosen to study the temperature dependence of the permeability. The oocyte volumetric responses during the experiments were recorded using DP71 CCD. The experimental videos were converted into images with a 10 s interval. The ImageJ software was used to analyze these images to determine the volume response of the oocytes. The oocytes were assumed to be spherical, and the volumes of the oocytes were calculated by processing the area of the oocytes in the image frames. We did not select any aspheric oocytes for practice. The oocyte volume data were normalized and substituted into Eqs. (2–4) (Supporting Information) to determine the 8


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membrane permeability coefficients. Fertilization of human oocytes and development of the fertilized oocytes. After blind screening huamn oocytes, to verify the outcome of the oocytes detection, we performed the insemination of oocytes by intracytoplasmic sperm injection (ICSI) using the patients' husband sperm. Fertilized oocytes were subsequently cultured individually in a droplet of cleavage medium (30 μl) under mineral oil (300-400 μl) at 37 °C with 6% CO2. Development of the fertilized oocytes was monitored for the formation of different stage embryos at various intervals about 5 days. Protocols of ICSI and embryo culture are detailed in the literature.40

 RESULTS AND DISCUSSOIN Characterization of solution concentration change in microperfusion chamber. For investigating the flow of CPA solution in microchannel, fluorescein disodium salt was used as a fluorescent tracer.41 As shown in Figure 2A, three typical positions (P1, P2 and P3) of the barrier in microchannel are chosen. The variation of CPA concentration from the inlet to the location of the oocyte can be characterized according to the corresponding fluorescence intensity. From Figure 2A, we can see the change of solution flow within the microchannel from a macroscopic level. As shown in Figure 2B, it indicates that the fluorescence intensity of the position P1 is a little bit greater than the position P2 and P3, due to faster to reach equilibrium than the other two positions. Therefore, it reveals that it is very scientific to use the arc shape design to assist the oocyte to stay in the middle of the channel. It can be seen that the solution concentration gradually increases along the channel while the continuous concentration change can be used to reduce osmotic shock. Thus, the microfluidic device is efficient and reliable. Moreover, we conducted the experiment with three different velocities (4, 8, 12 9


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μl/min) at P1, the equilibrium time of solution concentration is in corresponding with the flow rate (Figure 2C). The representative images of fluorescence intensity change at three typical positions (12 μl/min) are shown in Figure 2D.

Figure 2. Characterization of solution concentration change in the microfluidic channel. (A) The photographs of fluorescent solution flow in microchannel. P1-P3 are typical positions used to characterize solution flow. (B) Relative fluorescence intensity change of the three typical positions. (C) Relative fluorescence intensity change of three different flow rate at P1. (D) The fluorescent images of three typical positions in microchannel.

Simulation of CPA solution concentration profile in microperfusion chamber. To better understand the change of CPA solution concentration in microperfusion chamber, we used COMSOL 10


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Multiphysics® to build the model to simulate the CPA concentration change. The simulation of CPA concentration variation was divided into two kinds of solutions, 1 M EG and 1 M PG. The simulated CPA concentration change at the location for trapping oocytes are shown in Figure 3A&B. It can be seen that there is no significant difference between the two kinds of solutions, this should be due to the osmotic pressure of the two CPA solutions are almost equal. The typical images of two CPA concentration change in the microchannel at different time (25 °C) are also shown in Figure 3A&B. In Figure 3C&D, the front view and side view show the cross-sectional and longitudinal section of CPA (EG) solution concentration variation in the microchannel, respectively. It indicates that the CPA concentration gradually increases along the microchannel and the location for trapping oocyte is the fastest to achieve the final concentration. These results exhibit the flow change of the CPA solution in the microchannel and demonstrate that the microfluidic device is feasible.

Figure 3. Simulation of solution concentration profile in the microfluidic channel. (A) and (B) Solution 11


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concentration curves and flow changes of EG and PG throughout the whole microchannel, respectively. (C) and (D) The front and side view of CPA (EG) concentration change at P1, respectively.

Determination of inseminated-unfertilized oocyte membrane transport properties. To determine, if temperature affects the hydraulic conductivity and solute permeability of inseminated-unfertilized oocytes in different CPA solutions. We tested the oocytes at different temperatures and analyzed the experiment data. The Lp and Ps determined with two different CPA solutions at four experimental temperatures are shown in Table S1. These data indicate that the inseminated-unfertilized oocyte membrane permeability in the presence of EG is smaller than that in the presence of PG at the same temperature. The normalized oocyte volumetric change to the osmotic pressure change at four experimental temperatures are shown in Figure 4A&B. It can be observed that for the addition of either CPA, the oocyte volume experienced a shrinkage followed by a recovery, which is similar to the volume responses of most cells during permeable CPA loading reported in previous study.42 Meanwhile, we can see that the oocyte experiences a more severe volume shrinkage during loading EG than the loading PG at the same temperature, which implies that the permeability of EG is less than that of PG at that certain temperature. Compared our results with previously published results43 under similar experimental conditions, the measured Lp and Ps are slightly smaller than the previously published normal oocytes. This may indicate that the oocyte membrane permeability has a relationship with fertilization failure.

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Figure 4. Biophysical properties of human inseminated-unfertilized oocytes membrane. (A) and (B) Normalized oocyte volume changes of representative oocytes exposed in EG and PG at four temperatures and the corresponding curve fitting results (lines) with 2-P model. (C) and (D) Arrhenius plots of the temperature-dependent membrane permeability coefficients. (E) and (F) The values of Lp and Ps at four temperatures, respectively. (G) The activation energies for water and CPA transport across the oocytes membrane.

To verify their temperature dependence, the Lp and Ps measured in 1 M EG and 1 M PG solution were fitted to the Arrhenius relationship, as shown in Figure 4C&D, respectively. Based on Eq. (5) and Eq. (6) (Supporting Information), a least-square linear fit was applied to the experimental data. The oocyte membrane permeability coefficients for water (Lpg) and CPA (Pag) at the reference temperature (T0 = 273.15 K) and the activation energy for the water (ELp) and CPA (EPa) permeability coefficients 13


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are listed in Table S2 (from 25 °C to 0 °C). These parameters could be used for optimization of CPA loading/unloading by minimizing the osmotic injury. The Lp and Ps at four experimental temperatures, and activation energies are given in Figure 4E-G, respectively. At each of the different temperatures, there is no significant difference among the Lp or Ps values except for the value of Ps at 25 °C. Additionally，we observed either Lp or Ps has a dependency on CPA and both Lp and Ps increase with increasing temperature. For activation energy, the transport of PG is greater than that of EG. It indicates that the temperature dependence of the permeability coefficient is stronger in the presence of PG than EG. It also shows that the activation energy for the water transport is smaller than the CPA transport. Oocyte viability test post microperfusion. To investigate whether the procedure we used has an adverse impact on oocyte, the oocyte viability test was performed using acridine orange/ethidium bromide (AO/EB) standard Live/Dead Staining Kit (KeyGen BioTECH Co. Ltd. China). The isotonic solution containing oocyte and AO/EB (1:1 v/v) were mixed at a ratio of 25:1 (v/v) and incubated at room temperature for about 3 min. Then oocytes were injected into the perfusion channel and captured at specific location, and fluorescent images were taken before and after the perfusion experiments, respectively. Live oocytes stained in green and dead oocytes stained red. The representative photomicrographs of oocyte volume changes to the addition of PG and EG are shown in Figure 5A&B, respectively. From previous research, we know that the cell dehydration is caused by the efflux of water driven by differences in intracellular and extracellular chemical potentials, while the volume recovery is a consequence of the transport of water and CPA into cells.44 Comparing Figure 5A and 5B, given the osmolality of the 1 M EG and 1 M PG are almost the same, we can conclude that the oocyte dehydration rate and volume recovery decreased with decreasing temperature. The typical fluorescent images are 14


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shown in Figure 5C. It indicates that the oocytes are still alive pre-and post-microperfusion. This demonstrates that the microperfusion device is feasible and safe for oocyte. After the experiment, the oocytes can be retrieved for further research.

Figure 5. Typical photomicrographs of inseminated-unfertilized oocytes in microperfusion channel. (A) Volume response of a typical oocyte after being exposed to PG solution at 25 °C. (B) Volume response of a typical oocyte after being exposed to EG solution at 5 °C. (C) The typical fluorescent images of oocytes before and after perfusion at 0, 5, 15, 25 °C.

Mechanism of high-quality oocytes selection and in vitro development validation. Compared to normal oocytes,45 there is difference in membrane permeability parameters between the normal and inseminated-unfertilized oocytes. We utilized the two permeability parameters obtained at 25 °C to 15


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predict the oocyte volume change. The predicted curves are shown in Figure 6A, the green curve represents normal oocyte, and the red curve represents the inseminated-unfertilized oocyte (Among them, the dotted line is obtained by a 10% change in the solid line parameter). The gray dots represent the experimental data of inseminated-unfertilized oocyte. It shows that the normalized volume change curve has a certain difference between the two kind oocytes. Therefore, we deduce that the difference in oocyte membrane permeability can be used to detect high-quality oocyte. To verify this inference, the oocytes isolated from COCs were collected for blind screening. As shown in Figure 6A, the green and red circles represent the high-quality and poor-quality oocytes data obtained by experiment, respectively. It demonstrates that oocytes can be divided into two subpopulations based on the membrane permeability. High-quality oocytes are more active in physiological activities, as makes the oocyte more adaptable to the external environment and respond more quickly in volume change upon osmotic shift. Our result is essentially consistent with the previous finding that high-quality oocytes have better mechanical properties.23 To further verify the aforementioned conclusion, the experimental oocytes were retrieved for ICSI to test their developmental capacity. The results of the developmental potency are shown in Figure 6B. There is no significant difference in fertilization rate between the two groups, which may be caused by ICSI insemination. For the fertilized oocytes, the cleavage rate of selected high-quality oocyte group is much higher than the poor-quality oocyte group (100% vs. 33.3%). The subsequent blastocyst rate is 60% vs. 0% for high-quality and poor-quality oocyte group, respectively. The data is shown in Table S3. It is worth noting that the poor-quality oocytes can only develop into 4-cells stage. These results indicate that our method for detecting high-quality oocytes is feasible and effective. A typical photograph of the process of ICSI is shown in Figure 6C. And the micrographs showing the morphology of the oocytes, 16


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the fertilized oocytes at 2-PN, 4-cells, 8-cells and blastocyst stage are also shown in Figure 6C. Comparing to the high-quality oocyte, the poor-quality oocyte do not acquire 8-cells and subsequent embryo. According to the results, we can determine the oocyte status by experimentally observing the change of oocyte volume at a specific temperature, so as to screen out the high-quality oocytes. Therefore, we believe this method can significantly enhance the fertilization rate of oocyte and embryo quality in ART. Moreover, compared with the other researches, our method has the advantages of easy fabrication, low cost, simple operation, and so on. The advantages and disadvantages between our method and the existing methods are shown in Table 1. Table 1. Comparison between this study and the existing methods. Method

Cost

Detection speed

Accuracy

Device fabrication

Experimental procedure

Device reusability

Morphological methods5-7

Low-cost

Very fast

Very low

-

Simple

Yes

Optical methods18, 19

Expensive

Fast

Low

Difficult

Complex

Yes

Mechanical methods22, 26 Genetic screening methods27-29

Expensive

Fast

Medium

Difficult

Complex

Yes

High expensive

Slow

High

-

Complex

No

Our method

Low-cost

Fast

High

Easy

Simple

Yes

Note: Accuracy refers to developmental capacity of oocytes.

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Figure 6. The mechanism of high-quality oocytes selection and in vitro development verification. (A) The high-quality and poor-quality oocytes are distinguished by the difference of membrane permeability at the same condition. (B) Comparison of development capacity between screened high-quality oocyte and poor-quality oocyte. (C) Typical photomicrographs of ICSI and the morphology of the original mature oocytes and subsequent fertilized oocytes development (2-pronuclei, 4-cells, 8-cells and blastocyst embryos).

 CONCLUSIONS In this study, we propose a novel and effective approach of on-chip detection of high-quality human oocytes. This approach has the advantages of low cost, easy fabrication, high reusability and simple operation. By using a self-made microfluidic chip, we detected the membrane permeability of the oocytes. Experimental results indicate that oocytes can be divided into high-quality and poor-quality groups according to the difference in their membrane permeability. Subsequent IVF and development 18


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experiments found that there is a significant difference between the developmental capacity of the screened high-quality oocytes and poor-quality oocytes (the blastocyst rate is 60% vs. 0%). This indicates that our method can successfully detect high-quality oocytes and improve the ART outcome. In addition, this study enriched the important and fundamental data of oocyte membrane permeability under different conditions. We believe that this approach may find wide applications in the field of reproductive medicine, and will greatly promote the development of assisted reproductive medicine and the fertility preservation.



ASSOCIATED CONTENT

Supporting information The Supporting Information is available free of charge on the ACS Publications website at DOI: XXX. Theory of mass transport across cell membrane, the permeability parameters of oocytes, the developmental capacity of two kind oocytes. (PDF)

 AUTHOR INFORMATION Corresponding Author *Email: [email protected] (G.Z.); *Email: [email protected] (Y.C.).

ORCID Gang Zhao: 0000-0002-0201-1825 Zhongrong Chen: 0000-0002-4007-4800 Kashan Memon: 0000-0003-0339-6007 19


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Fazil Panhwar: 0000-0003-0251-6935 Meng Wang: 0000-0003-1676-8172

Author Contributions # Equal

contribution.

Notes The authors have no competing interests to declare.

 ACKNOWLEDGMENTS This work was supported by the National Natural Science Foundation of China (Grant nos. 51476160 and 11627803). This work was partially performed at the USTC Center for Micro- and Nanoscale Research and Fabrication.

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For TOC only

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Figure 1. Schematic illustration of the microfluidic perfusion system. (A) The composition of the whole system. (B) The 2D view of the microfluidic perfusion device and experimental principle. (C) The 3Ddisassembled view of the microfluidic perfusion device. Note: drawing not to scale. 124x97mm (300 x 300 DPI)


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Figure 2. Characterization of solution concentration change in the microfluidic channel. (A) The photographs of fluorescent solution flow in microchannel. P1-P3 are typical positions used to characterize solution flow. (B) Relative fluorescence intensity change of the three typical positions. (C) Relative fluorescence intensity change of three different flow rate at P1. (D) The fluorescent images of three typical positions in microchannel. 109x118mm (300 x 300 DPI)


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Figure 3. Simulation of solution concentration profile in the microfluidic channel. (A) and (B) Solution concentration curves and flow changes of EG and PG throughout the whole microchannel, respectively. (C) and (D) The front and side view of CPA (EG) concentration change at P1, respectively. 80x101mm (300 x 300 DPI)


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Figure 4. Biophysical properties of human inseminated-unfertilized oocytes membrane. (A) and (B) Normalized oocyte volume changes of representative oocytes exposed in EG and PG at four temperatures and the corresponding curve fitting results (lines) with 2-P model. (C) and (D) Arrhenius plots of the temperature-dependent membrane permeability coefficients. (E) and (F) The values of Lp and Ps at four temperatures, respectively. (G) The activation energies for water and CPA transport across the oocytes membrane. 82x101mm (300 x 300 DPI)


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Figure 5. Typical photomicrographs of inseminated-unfertilized oocytes in microperfusion channel. (A) Volume response of a typical oocyte after being exposed to PG solution at 25 °C. (B) Volume response of a typical oocyte after being exposed to EG solution at 5 °C. (C) The typical fluorescent images of oocytes before and after perfusion at 0, 5, 15, 25 °C. 82x112mm (300 x 300 DPI)


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Figure 6. The mechanism of high-quality oocytes selection and in vitro development verification. (A) The high-quality and poor-quality oocytes are distinguished by the difference of membrane permeability at the same condition. (B) Comparison of development capacity between screened high-quality oocyte and poorquality oocyte. (C) Typical photomicrographs of ICSI and the morphology of the original mature oocytes and subsequent fertilized oocytes development (2-pronuclei, 4-cells, 8-cells and blastocyst embryos). 174x92mm (300 x 300 DPI)


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Sensing cell membrane biophysical properties for ... - ACS Publications

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