Printer Friendly

Integration of sperm motility and chemotaxis screening with a microchannel-based device.

Since the first "test-tube baby" in 1978 (1), in vitro fertilization (IVF) [6] has been widely used in cases of infertility problems. In recent years, the success rate and safety level have been enhanced by the development of new technologies such as embryo freezing (2), preimplantation genetic diagnosis (3), and intracytoplasmic sperm injection (ICSI) (4). Sperm screening is an important step for the success of fertilization (5). Currently, sperms are clinically selected in test tubes on the basis of motility rather than degree of health. Under physiological conditions, different mechanisms have been shown to guide sperms successfully along the female genital tract. In mammals, these mechanisms are thought to be a combination of chemotaxis, thermotaxis, and probably oviductal contractions (6).

Motility is regarded as one of the most important parameters related to sperm quality, and asthenospermia is a common reason for male infertility. Procedures that could safely and efficiently select motile sperms would be desirable. Currently, the swim-up method and the density-gradient centrifugal method are used for this purpose (7).

Chemotaxis, the movement of cells toward a concentration gradient of chemoattractant, was first discovered as a sperm guidance mechanism in marine species (8). In the last decade, the mechanism of sperm chemotaxis in mammals has been extensively debated. The technical difficulty lies in the low signal-to-noise ratio obtained in the chemotaxis assay and suboptimal assays that cannot distinguish real chemotaxis from sperm accumulation caused by other factors (9). Recent advances demonstrate that the low signal-to-noise ratio is because only capacitated spermatozoa are chemotactically responsive and the percentage of capacitation is low (approximately 10% in humans) (10). Sperm chemotaxis has been demonstrated in frogs, mice, rabbits, and humans, and potential attractants include heat-stable peptides and progesterone in follicular fluid (11).


The manipulation and analyses of biomolecules, cells, and even organisms have been recently made possible using lab-on-a-chip devices (12-14). Microfluidics has shown potential in assisted reproductive technology (ART) because of its rapid analysis capability on small sample volumes and the potential to reduce manual operation (15-17). Several microfluidic devices have been developed for sperm testing (15, 16). Kricka and colleagues (15, 16) developed a motile sperm-testing microchip with tortuous channels. A microfluidic device based on sperm-sorting method was proposed by Cho et al. in 2003 (17) that used a parallel laminar flow stream to distinguish motile from poor-quality sperms. Koyama et al. (18) were the first to use microfluidics for sperm chemotaxis testing via a 3-channel structure designed to form a spatially and temporally stable chemical gradient.

In this study, we describe the development of a microfluidic system that can sort healthy sperms on the basis of not only motility, but also chemotactic responsiveness. We sorted sperms with good motility using a width- and length-optimized microchannel, followed by a bibranch channel where cumulus cells were cultured in 1 branch to form a chemoattractant gradient to select sperms with good chemoattractive ability.

Materials and Methods


We obtained human tubal fluid (HTF) and potassium simplex optimized medium (KSOM) from Chemicon; hyaluronidase, pregnant mare serum gonadotropin, and human chorionic gonadotropin (hCG) from Sigma; fetal bovine serum (FBS) from PAA Laboratories; polydimethylsiloxane (PDMS, Sylgard 184) from Dow Corning Corp; and propidium iodide (PI), SYBR 14, and JC1 from Invitrogen.

Sexually mature (>8-week-old) ICR mice, raised in the Institute of Genetics and Developmental Biology of the Chinese Academy of Sciences, were housed in the Animal House of Tsinghua University and used in the experiments.


To facilitate the sperm motility screening, we optimized the width and length of the microchannel. To determine the channel width, the length was set as 7 mm and the widths varied as follows: 200 [micro]m, 500 [micro]m, 1 mm, and 1.5 mm. For the channel length, the width was set as 1 mm and the lengths tested were 5 mm, 7 mm, 10 mm, and 15 mm. The depth of the channel was set at 25 [micro]m to facilitate the in-focus observation of sperms swimming from inlet to outlet and guarantee the accuracy of sperm counting. The radii of the inlet and outlet pools were 2 mm and 1.25 mm, respectively. Fig. 1A shows the structure of the microfluidic chip for motility screening.

The sperm motility and chemotaxis testing structure, imitating the female genital tract (Fig. 2A), comprised 4 divisions: (1) the inlet pool, where sperms can be added and start to swim, (2) a straight channel with optimized parameters to screen for sperms with good motility, (3) a round chamber where the motile sperms can be collected and a 2-dimensional chemical gradient can be generated, and (4) symmetrical bibranch channels with 2 outlet pools used as chemoattractant sources. The 2 branches form a 90-degree angle and are both 5 mm in length and 1 mm in width. The radius of the inlet and outlet pools was set at 2 mm, which is the same as the inlet pool of the motility-screening channel. The radius of the round chamber was kept at 1.25 mm, the same as the outlet pool of the motility-screening channel. Fig. 2B shows the schematic diagram for the bibranch microchannel enabling both sperm motility and chemotaxis evaluation.


We constructed microfluidic chips by use of standard photolithography and micromolding procedures (19). Briefly, the SU-8 photoresist was patterned onto a 4-inch silicon wafer to form a master. Liquid PDMS prepolymer was poured on the master mold, cured at 70 [degrees]C for 1.5 h, and then peeled off the master mold, producing the final replica with the channel structures. The PDMS replica was then diced and plasma-bonded to a glass slide. Figs. 1B and 2C show the device with only the motility-screening channel and with both the motility and chemotaxis testing channels, respectively.


Eight-week-old female ICR mice were superovulated by giving an intraperitoneal (ip) injection of 10 IU pregnant mare serum gonadotropin 62 h before collection, followed by an ip injection of 10 IU hCG 14 h before collection. Mice were killed by cervical dislocation, and we collected the cumulus-oocyte-complexes (COCs) from the oviducts in HTF medium. We used 3-minute digestion with 3% hyaluronidase at 37 [degrees]C to separate primary cumulus cells from oocytes. We then added FBS to a final concentration of 10% to terminate the digestion. The cumulus cells were then spun down at 200g for 5 min and resuspended with HTF containing 10% FBS. To form the chemoattractant gradient, we seeded the homogeneous cumulus cell suspension to 1 of the outlet pools of the bibranch, and cells adhered 5-6 h later. Cells were usually planted at 60% confluency (approximately 1 x [10.sup.4] cells) and were ready for use after 24 h of culture. It is important to avoid turbulence of the fluid while planting the cells. Fig. 2D shows the normally growing cumulus cells cultured in 1 of the outlet pools 24 h after seeding (bright field, 200X).


An 8-week old male ICR mouse was killed by cervical dislocation. The cauda epididymides and vasa deferentia from 2 sides of the mouse were immediately dissected, placed in 250 [micro]L HTF medium preequilibrated overnight. Each cauda epididymis was then cut into 5-7 pieces with ophthalmology scissors to allow motile sperms to swim out. We placed the sperm suspension in a standard humidified 5% C[O.sub.2] incubator at 37 [degrees]C for 30 min for capacitation and assessed sperm concentration by use of a hemocytometer (Qiujing).

To assess motility, we added approximately 25 000 sperms were added to the inlet pool of the sperm motility-screening device. After 5-10 min of swimming, sperms started to accumulate in the outlet pool consistently in each microchannel. After 10 min of swimming, 5-s video recordings were captured at the central region of the inlet pool and near the entrance of the outlet pool. We used a DP-71 charge-coupled device (CCD) (Olympus) coupled with an inverted microscope (DM-IRB, Leica) for video recording (200 x); the visual field was randomly selected as a representative region for the inlet or outlet pool.

For the sperm motility and chemotaxis testing device, we added approximately 25 000 sperms to the inlet pool. After 10 min of swimming, sperms started to reach the bifurcation consistently. A 15-min video recording captured sperms heading toward different branches. Two experimental groups were set up with cumulus cells planted in either pool A or pool B. More than 40 sperms were recorded for each experiment. The videos were carefully viewed to count the number of sperms passing through [L.sub.1] or [L.sub.2], respectively (Fig. 2E). Control experiments were set up in the same manner, except that in each experiment, cumulus cells were planted in both or neither of the 2 pools. We replicated the experiments independently 3 times. We used a DP-71 CCD coupled with an inverted microscope for video capture (50X). (See accompanying supplemental video at 56/8/1270 and select Video.)


We used the Live/Dead Sperm Viability Kit (Invitrogen), which uses a mixture of SYBR 14 dye and PI, to test the intact membrane rate of the sperm sample (20). We added approximately 25 000 sperms to the inlet pool of the sperm motility screening device, which was prefilled with HTF containing 1 [micro]mol/L SYBR 14 and 20 [micro]mol/L PI. After 10-min incubation at 37 [degrees]C in 5% C[O.sub.2], the sperms with intact membranes stained green and the ones with damaged membranes stained red. The stained sperms (approximately 10 [micro]L) in the inlet or outlet pool were mounted on the glass slide and examined under a fluorescent microscope (Leica DMIRB, 400 x; the excitation wavelengths of SYBR 14 and PI are 488 and 535 nm, respectively; emission wavelengths of these 2 dyes are 518 and 617 nm, respectively).

The first J-aggregate-forming cationic dye (JC-1) is a cationic dye that indicates mitochondrial membrane potential. The decrease of red/green fluorescence intensity ratio indicates depolarization of mitochondria, which negatively correlates with cell viability (21). The shift is due to concentration-dependent formation of J-aggregates that fluoresce red. We added approximately 25 000 sperms to the inlet pool of the sperm motility screening device, which was prefilled with HTF containing 10 mg/L JC-1, and incubated at 37 [degrees]C for 15 min. The stained sperms (approximately 10 [micro]L) in the inlet or outlet pool were mounted on the glass slide and examined under a fluorescent microscope (400x). The excitation wavelengths of the monomer form and J-aggregate form are 514 and 585 nm; emission wavelengths are 529 and 590 nm, respectively.


For the sperm motility screening experiment, we captured 5-s video recordings (200X) at both inlet pool and outlet pool for each microchannel. Motile (progressively motile plus nonprogressively motile) and total sperm number were counted in the randomly selected field for the inlet and outlet pools, respectively. Two indices were adopted in the experiment. One is sperm motility in the outlet pool, defined as the percentage of motile sperms in the total sperm population. The other is the relative sperm count in the outlet pool, defined as the sperm number in the selected field in the outlet pool normalized to the sperm number in the selected field in the inlet pool. The sperm motility and count in the outlet pool were compared among groups with different channel dimensions. For comparisons, we used 1-way ANOVA (22) and appropriate post hoc testing if differences were significant (P < 0.05).

For convenience in evaluating sperm chemotaxis in the current device, we derived a parameter called "chemotaxis index" to assess the characteristics of sperm chemotaxis, represented as the ratio of the number of sperm swimming toward pool A vs the number of sperm swimming toward pool B. We performed the Student t-test (paired, 2-tail) to assess the chemotactic response difference between the experimental and control groups. Groups 1 and 2 were combined as an experimental group by averaging the chemotaxis index of group 1 and the reciprocal of the chemotaxis index of group 2. Three replicates were performed, and a probability of P < 0.05 was considered statistically significant (23). All computations were made using SPSS software (SPSS Inc.).



We adopted 2 indices to determine the channel parameters--1 was sperm motility and the other was the relative sperm count in the outlet pool. Sperm motility in the inlet pool was approximately 60% (mean for all the microchannels) and reached approximately 85% in the outlet pool, which represents 20%-30% more than in the inlet.

For the width optimization, we saw no statistically significant difference among the various width channels for either sperm motility (Fig. 3A) or relative sperm count (data not shown) in the outlet pool. Among the channels of 1-mm width by 5- to 15-mm length, we found that there was a significant difference for both sperm motility and relative sperm count between the 4 groups (P = 0.022 and 0.011, respectively, 1-way ANOVA). For further analysis, we carried out multiple comparisons between each pair of groups. With regard to sperm motility, the mean motility in the outlet of 5-mm channel was significantly lower than the other 3 groups (least significant difference test, P = 0.041, 0.011, and 0.005 for 5-mm group vs 7-, 10-, and 15-mm groups, respectively), whereas no significant difference was observed among the other 3 groups. As for the relative sperm count, the mean values for the 5and 7-mm groups, but not the 10-mm group, were significantly higher than that of the 15-mm group (least significant difference test, P = 0.0026, 0.014, and >0.1 for 15-mm group vs 5-, 7-, and 10-mm groups, respectively). Taking both indices into account, we considered a 7-mm channel as the 1 with best performance [sperm motility 82.6% (2.9%); relative sperm count 36.3% (8.3%)] and used this length in subsequent experiments.


The sperms stained green in the total population were increased in the outlet compared to the inlet pool (Fig. 4A and B). The fluorescence result of JC-1 staining is shown in Fig. 4C and D. The higher red-to-green ratio in outlet pool further confirmed the effectiveness of the optimized channel in sperm-motility evaluation.


For the purpose of investigating the chemotactic behavior of the sperms, we proposed a new assay using microchannel-cultured cumulus cells as the chemoattractant source to achieve greater stability and a closer match to the in vivo environment.

To establish a stable and objective method for the evaluation of chemotaxis, we set up 4 groups of sperm passing through microchannels with different chemoattractant patterns. Group 1 had the cumulus cells planted only in outlet pool A; group 2 had the cumulus cells planted only in outlet pool B; group 3 had cumulus cells planted in both pools A and B; group 4 had no cumulus cells planted in any pool. Therefore, if sperm chemotaxis was taking place, the count of sperm swimming toward pool A would be expected to be relatively high in group 1 and low in group 2, and we would expect the chemotaxis index to be > 1 for group 1 and < 1 for group 2. For groups 3 and 4, we would expect the chemotaxis index to be approximately 1, since there was no chemical gradient under these 2 situations and sperm would show no bias of swimming toward either pool A or B.

Fig. 5 illustrates that the sperm were chemotactically attracted to cumulus cells. The chemotaxis indices were 1.239, 0.815, 1.061, and 0.963, respectively, for each experimental group and demonstrated a significant fraction of sperm traveling toward the cumulus cells compared with the control (combined group 1 and 2 as experimental group, P = 0.0022 vs group 3 and P = 0.039 vs group 4). The subpopulation of chemotactically responsive sperms was defined as the difference between sperms swimming toward the cumulus cells and sperms swimming toward the opposite branch. This mean percentage was 10.2% (n = 6, SD 5.2%), consistent with previous studies showing only a small fraction of total sperms being chemotactic (approximately 10%) (18,24). After screening, the percentage of chemotactically responsive sperms increased to 18.6% (proportion of 10.2% chemotactically responsive sperms in a total of 55.1% selected sperms).



We describe the development of an efficient microfluidic system by incorporating both motility and chemotactic responsiveness for sorting sperms in vitro. Among the different approaches used for motility screening, a method using microchannels surpasses others in simplicity, user friendliness, and resemblance to the physiological environment. The whole assay was performed automatically in stable fluid and thus avoided the mechanical force involved in conventional sperm selection procedures that may cause DNA fragmentation (25, 26). The material used was PDMS, which is biologically compatible with a variety of cell types, and the screening channel was formed between PDMS and the glass slide.

Our first goal was to study how variation in microchannel width and length altered in vitro motile sperm selection. When we examined a width range of 200 [micro]m to 1.5 mm, we saw no significant difference between the various microchannels. When we examined a length range of 5-15 mm, however, short channels (5 mm) could not properly distinguish motile sperms from sperms with no or low motility, and increasing the length of the channel increased the percentage of motile sperms. The length of microchannel beyond the optimal value, however, dramatically decreased the sperms that could reach the outlet. This is expected, as the long distance could exhaust the sperms and thus may not be suitable for downstream fertilization applications. Although the study was done using murine sperms, we propose that the system could also be applied to other species with readjustment of the above parameters. In our current study, sperms were allowed to swim for only 10 min before data collection, and the channel depth was set at 25 [micro]m to satisfy the in-focus observation. The time point set for data collection and the depth of the testing channel to some extent limited the sperm amount obtained in the outlet. However, the time of sperm screening and the channel depth could be increased to further enrich the sperms in the outlet.


The efficiency of the microchannel was further confirmed by fluorescent staining. The percentage of motile sperms in the outlet pool was increased compared with the inlet pool. Another important finding was that after swimming through the channel, the phenomenon of sperm "groups" or "trains" was largely eliminated. It is common for rodent sperms to form sperm groups by use of the apical hook in their head that might help to increase swimming velocity and thrusting force in vivo (27); however, this may not be suitable for in vitro fertilization. One reason is that for IVF applications, sperm competition is not as important as in vivo fertilization, which needs sperm groups to surpass others to reach the egg. In addition, because the mechanisms for dispersal of aggregated sperms before fertilization, involving in part acrosome reaction, are missing in vitro (28), sperm groups may even hamper subsequent fertilization. In summary, sperms after screening not only had excellent motility scores, but also detached from each other, potentially further increasing the efficiency for IVF.


Currently, 3 types of assays are used for studying sperm chemotaxis in mammals. The first is an accumulation assay in which sperms sense the chemoattractant gradient and accumulate near or at its source. The second is the so-called choice assay, in which sperms choose between 2 or more wells containing control or attractant buffer. The third is to directly trace video-recorded tracks made by sperms sensing the chemoattractant (9). Our assay could theoretically be classified as the choice assay and can efficiently differentiate the chemotaxis from sperm accumulation or trapping.

The major difference of our approach compared with previous assays is that we used in vitro cultured cumulus cells as the chemoattractant source instead of follicular fluid or progesterone-containing medium. Prior studies using follicular fluid failed to generate consistent results (29). Much effort has been expended to clarify the working concentration of follicular fluid (30). It is important to bear in mind, however, that chemotaxis to the follicular fluid is unlikely to occur in vivo because it is only released as a single event at ovulation and cannot be maintained throughout the life of the egg in the female genital tract (31). With our proposed method, we can bypass this issue and provide a method that better mimics the in vivo environment. It is believed that the oocytes as well as the surrounding cumulus cells can secrete components that guide sperms to the oocytes (31). These components are consistently secreted after ovulation, of which progesterone is the major chemoattractant (32). We planted the cumulus cells in the microchannels 24 h before performing the assay to generate a chemoattractant gradient. Based on our observation, cumulus cells could adhere to the substrate 5-6 h after seeding and spread effectively after overnight culture. They began to show changes in morphology 48 h after seeding, without a medium change. The device we designed also mimics the in vivo structure: the screening channel resembles the vagina; the diffusion chamber could store and buffer the sperms just like a female uterus; the bibranch channel functions as the oviducts; and the in vitro cultured cumulus cells provide the chemoattractive function of the COCs in vivo. We found a moderate but significant chemotaxis phenomenon that sperms tended to swim to the wells containing cumulus cells. The total percentage of chemotactic responsive sperms was 10.2%, consistent with previous studies reporting values ranging from 7% to 13% in mice (18, 24). More studies are needed to elucidate the progesterone distribution in this system and the mechanism of how cumulus cells attract sperms.

Our microchannel-based device allows motility screening and chemotaxis testing simultaneously, demonstrating its potential for clinical treatment of infertility. Further work would be the building of a labon-a-chip type system that could integrate sample preparation, biological or chemical reaction, and data analysis in miniaturized channels or chambers for assisted reproduction applications. Such a system could surpass the conventional methods, as it could be fully automated and efficient and could closely resemble physiological conditions.

Author Contributions: All authors confirmed they have contributed to the intellectual content of this paper and have met the following 3 requirements: (a) significant contributions to the conception and design, acquisition of data, or analysis and interpretation of data; (b) drafting or revising the article for intellectual content; and (c) final approval of the published article.

Authors' Disclosures of Potential Conflicts of Interest: No authors declared any potential conflicts of interest.

Role of Sponsor: The funding organizations played no role in the design of study, choice of enrolled patients, review and interpretation of data, or preparation or approval of manuscript.


(1.) Steptoe PC, Edwards RG. Birth after the reimplantation of a human embryo. Lancet 1978;2:366.

(2.) Trounson A, Mohr L. Human pregnancy following cryopreservation, thawing and transfer of an eight-cell embryo. Nature 1983;305:707-9.

(3.) Handyside AH, Kontogianni EH, Hardy K, Winston RM. Pregnancies from biopsied human preimplantation embryos sexed by Y-specific DNA amplification. Nature 1990;344:768-70.

(4.) Palermo G, Joris H, Devroey P, Van Steirteghem AC. Pregnancies after intracytoplasmic injection of single spermatozoon into an oocyte. Lancet 1992;340:17-8.

(5.) Parrish JJ, Krogenaes A, Susko-Parrish JL. Effect of bovine sperm separation by either swim-up or Percoll method on success of in vitro fertilization and early embryonic development. Theriogenology 1995; 44:859-69.

(6.) Eisenbach M, Giojalas LC. Sperm guidance in mammals: an unpaved road to the egg. Nat Rev Mol Cell Biol 2006;7:276-85.

(7.) Xu L, Lu RK, Chen L, Zheng YL. Comparative study on efficacy of three sperm-separation techniques. Asian J Androl 2000;2:131-4.

(8.) Miller RL. Chemotaxis during fertilization in the hydroid Campanularia. J Exp Zool 1966;162:23-44.

(9.) Eisenbach M. Sperm chemotaxis. Rev Reprod 1999;4:56-66.

(10.) Cohen-Dayag A, Tur-Kaspa I, Dor J, Mashiach S, Eisenbach M. Sperm capacitation in humans is transient and correlates with chemotactic responsiveness to follicular factors. Proc Natl Acad Sci U S A 1995;92:11039-43.

(11.) Villanueva-Diaz C, Arias-Martinez J, Bermejo-Martinez L, Vadillo-Ortega F. Progesterone induces human sperm chemotaxis. Fertil Steril 1995;64:1183-8.

(12.) Cheng J, Sheldon EL, Wu L, Uribe A, Gerrue LO, Carrino J, et al. Preparation and hybridization analysis of DNA/RNA from E. coli on microfabricated bioelectronic chips. Nat Biotechnol 1998; 16:541-6.

(13.) Wang L, Zhu J, Deng C, Xing WL, Cheng J. An automatic and quantitative on-chip cell migration assay using self-assembled monolayers combined with real-time cellular impedance sensing. Lab Chip 2008;8:872-8.

(14.) Shi W, Qin J, Ye N, Lin B. Droplet-based microfluidic system for individual Caenorhabditis elegans assay. Lab Chip 2008;8:1432-5.

(15.) Kricka LJ, Nozaki O, Heyner S, Garside WT, Wilding P. Applications of a microfabricated device for evaluating sperm function. Clin Chem 1993;39: 1944-7.

(16.) Kricka LJ, Faro I, Heyner S, Garside WT, Fitzpatrick G, McKinnon G, et al. Micromachined analytical devices: microchips for semen testing. J Pharm Biomed Anal 1997;15:1443-7.

(17.) Cho BS, Schuster TG, Zhu X, Chang D, Smith GD, Takayama S. Passively driven integrated microfluidic system for separation of motile sperm. Anal Chem 2003;75:1671-5.

(18.) Koyama S, Amarie D, Soini HA, Novotny MV, Jacobson SC. Chemotaxis assays of mouse sperm on microfluidic devices. Anal Chem 2006;78: 3354-9.

(19.) Wang L, Wang H, Mitchelson K, Yu Z, Cheng J. Analysis of the sensitivity and frequency characteristics of coplanar electrical cell-substrate impedance sensors. Biosens Bioelectron 2008;24: 14-21.

(20.) Ericsson SA, Garner DL, Thomas CA, Downing TW, Marshall CE. Interrelationships among fluorometric analyses of spermatozoal function, classical semen quality parameters and the fertility of frozen-thawed bovine spermatozoa. Theriogenology 1993;39:1009-24.

(21.) Gravance CG, Garner DL, Baumber J, Ball BA. Assessment of equine sperm mitochondrial function using JC-1. Theriogenology 2000;53:1691-703.

(22.) Boiculese LV, Azoicai D, Rezus E, Rezus C, Dorneanu O. Comparing population means using the ANOVA method [in Romanian]. Rev Med Chir Soc Med Nat Iasi 2003;107:906-12.

(23.) Witt PL, McGrain P. Comparing two sample means t tests. Phys Ther 1985;65:1730-3.

(24.) Oliveira RG, Tomasi L, Rovasio RA, Giojalas LC. Increased velocity and induction of chemotactic response in mouse spermatozoa by follicular and oviductal fluids. J Reprod Fertil 1999;115:23-7.

(25.) Zini A, Finelli A, Phang D, Jarvi K. Influence of semen processing technique on human sperm DNA integrity. Urology 2000;56:1081-4.

(26.) Younglai EV, Holt D, Brown P, Jurisicova A, Casper RF. Sperm swim-up techniques and DNA fragmentation. Hum Reprod 2001;16:1950-3.

(27.) Immler S, Moore HD, Breed WG, Birkhead TR. By hook or by crook? Morphometry, competition and cooperation in rodent sperm. PLoS One 2007;2: e170.

(28.) Moore H, Dvorakova K, Jenkins N, Breed W. Exceptional sperm cooperation in the wood mouse. Nature 2002;418:174-7.

(29.) Eisenbach M. Mammalian sperm chemotaxis and its association with capacitation. Dev Genet 1999;25:87-94.

(30.) Eisenbach M, Tur-Kaspa I. Do human eggs attract spermatozoa? Bioessays 1999;21:203-10.

(31.) Sun F, Bahat A, Gakamsky A, Girsh E, Katz N, Giojalas LC, et al. Human sperm chemotaxis: both the oocyte and its surrounding cumulus cells secrete sperm chemoattractants. Hum Reprod 2005;20:761-7.

(32.) Oren-Benaroya R, Orvieto R, Gakamsky A, Pinchasov M, Eisenbach M. The sperm chemoattractant secreted from human cumulus cells is progesterone. Hum Reprod 2008;23:2339-45.

Lan Xie, [1,2], ([dagger]) Rui Ma, [1,2], ([dagger]) Chao Han, [1,2] Kai Su, [3] Qiufang Zhang, [4] Tian Qiu, [1,2] Lei Wang, [2] Guoliang Huang, [1,2] Jie Qiao, [4] Jundong Wang, [3] * and Jing Cheng [1,2,5] *

[1] Medical Systems Biology Research Center, Department of Biomedical Engineering, Tsinghua University School of Medicine, Beijing, China; [2] National Engineering Research Center for Beijing Biochip Technology, Beijing, China; [3] Shanxi Key Laboratory of Ecological Animal Science and Environmental Medicine, Shanxi Agricultural University, Taigu, Shanxi, China; [4] Center of Reproduction Medicine, Department of Obstetrics and Gynecology, Third Hospital of Peking University, Beijing, China; [5] The State Key Laboratory of Biomembrane and Membrane Biotechnology, Tsinghua University, Beijing, China.

[6] Nonstandard abbreviations: IVF, in vitro fertilization; ICSI, intracytoplasmic sperm injection; ART, assisted reproductive technology; HTF, human tubal fluid; KSOM, potassium simplex optimized medium; hCG, human chorionic gonadotropin; FBS, fetal bovine serum; PDMS, polydimethylsiloxane; PI, propidium iodide; ip, intraperitoneal; COC, cumulus-oocyte complex; CCD, charge-coupled device; JC-1, first J-aggregate-forming cationic dye.

([dagger]) L. Xie and R. Ma contributed equally to this work.

* Address correspondence to these authors at: Jundong Wang, Shanxi Key Laboratory of Ecological Animal Science and Environmental Medicine, Shanxi Agricultural University, Taigu, 030801, Shanxi, People's Republic of China. Fax +86-354-6222942; e-mail; Jing Cheng, Medical Systems Biology Research Center, Tsinghua University School of Medicine, Beijing 100084, The People's Republic of China. Fax +86-10-62773059; e-mail

Received March 17, 2010; accepted May 13, 2010.

Previously published online at DOI: 10.1373/clinchem.2010.146902
COPYRIGHT 2010 American Association for Clinical Chemistry, Inc.
No portion of this article can be reproduced without the express written permission from the copyright holder.
Copyright 2010 Gale, Cengage Learning. All rights reserved.

Article Details
Printer friendly Cite/link Email Feedback
Title Annotation:Automation and Analytical Techniques
Author:Xie, Lan; Ma, Rui; Han, Chao; Su, Kai; Zhang, Qiufang; Qiu, Tian; Wang, Lei; Huang, Guoliang; Qiao,
Publication:Clinical Chemistry
Date:Aug 1, 2010
Previous Article:Disposition of cannabinoids in oral fluid after controlled around-the-clock oral THC administration.
Next Article:Analysis of the size distributions of fetal and maternal cell-free DNA by paired-end sequencing.

Terms of use | Copyright © 2018 Farlex, Inc. | Feedback | For webmasters