Sperm Assessment Using Flow Cytometry
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State of the art in sperm assessment using flow cytometry
Flow cytometry is emerging as an important tool in the field of modern andrology for routine analysis of spermatozoa. Recently, application of flow cytometry in the artificial insemination industry especially for pig is a new approach. Until very recent, semen sample analysis was routinely performed by microscopical evaluation and manual techniques by laboratory operators; the analysis is affected by a wide imprecision related to variability among observers, influencing its clinical validity. The last decade, several new flow cytometric techniques have been introduced for farm animal semen assessment that enable a more detailed evaluation of several sperm characteristics. Here in this paper, an initiative has been taken to focus on a number of recent flow cytometry developments important for addressing questions in andrological tests.
After the invention of flow cytometry, sperm evaluation by traditional microscopic means became questioned due to the robust advantages of flow cytometry over the microscopic method. Due to the recent development of large number of fluroscence probes, flow cytometry is now capable of analyzing number of sperm characteristics like viability, capacitation, acrosomal integrity, membrane permeability, membrane integrity, mitochondrial status, DNA integrity, decondensation of DNA and differences between gamets based on sex. The application of flow cytometry to their detection allows increased numbers of spermatozoa to be assessed over a short time-period, provides the possibility of working with small sample sizes, increases the repeatability of assessment, removes the subjectivity of assessment and allows simultaneous assessment of multiple fluorochromes. Flow cytometry is a technique capable of generating significantly novel data and allows the design and execution of experiments that are not possible with any other technique. Nowadays, semen evaluation using laboratory assays is extremely important to the artificial insemination industry to provide the most desired quality product to customers.
Future development of flow cytometric techniques will permit further advances both in our knowledge and in the improvement of assisted reproduction techniques. In this paper, the main semen parameters that can be analyzed with fluorochromes and adapted for use with a flow cytometer will be reviewed and the relationship of these tests to fertility will be discussed.
Semen evaluation is the single most important laboratory test that has helped us to identify clear-cut cases of fertility (Jarow et al., 2002), infertility or even of potential sub-fertility (Rodríguez-Martínez, 2007). Determination of the potential fertility of semen sample and, in the long run, of the male from which it has been collected is the ultimate goal of semen evaluations in clinically healthy sires. Methods are available that can sometimes estimate the potential fertilizing capacity of a semen sample and, in some cases, of the male (reviewed by Dziuk 1996; Rodríguez-Martínez et al. 1997a; Rodríguez-Martínez and Larsson 1998; Saacke et al. 1998; Larsson and Rodríguez-Martínez 2000; Rodríguez- Martínez 2000, 2003; Popwell and Flowers 2004; Graham and Mocé 2005; Gillan et al. 2005). The methods routinely used for evaluation of the quality of a semen sample involved an evaluation of general appearance (i.e. colour, contamination, etc.), volume, pH, sperm concentration, viability, morphology and motility. Most of these techniques are microscopic analyses that only measure a small number of spermatozoa within a population, are time-consuming, can be subjective and generally measure sperm attributes individually. Recently, limitations of semen evaluation methodology have been brought into sharp focus by controversies raised in the epidemiological literature. It should also be noted that such conventional measurements are prone to extreme inter-ejaculate variation, even when the laboratory methodology has been standardized. In the wake of this information, new opportunities have arisen for the development of methods for the diagnosis of male infertility, many of which have been shown to exhibit a prognostic value that eludes conventional semen profiling. Moreover, ejaculated spermatozoa are nowadays handled for use in assisted reproductive technologies, such as the artificial insemination of chilled, frozen-thawed or sexed semen, and IVF. Such handling implies semen extension, fluorophore loading, ultraviolet and laser illumination, high-speed sorting, cooling and cryopreservation, procedures that impose different degrees of change in sperm function following damage to sperm membranes, organelles or the DNA. Therefore, although several assays have been developed to monitor these sperm parameters, recently it is being claimed that buck of these procedures are incomplete, time consuming and laborious.
Flow cytometry in different technical applications offers many advantages for the analysis of sperm quality. Flow cytometry allows the simultaneous measurement of multiple fluorescences and light scatter induced by illumination of single cell or microscopic particles in suspension, as they flow very rapidly through a sensing area. The increasing use over the past decade of flow cytometry in the leading laboratories in human and veterinary andrology has dramatically increased our knowledge of sperm function under physiological and biotechnological conditions. Flow cytometers can acquire data on several subpopulations within a sample in a few minutes, making it ideal for assessment of heterogenous populations in semen sample. Initially developed in the 1960's, flow cytometry made automated separation of cells based on the unique recognition of cellular patterns within a population feasible (Hulett et al., 1969). Using such a separation approach, cellular patterns can be identified by assessing, in individual cells within a population, protein expression using fluorescently labeled antibodies and other fluorescent probes (Baumgarth and Roederer, 2000; Herzenberg et al., 2006).
Flow cytometry was first developed for medical and clinical applications such as haematology and oncology. These areas still account for the vast majority of publications on this technique, but during the past few years it has been used in other areas, such as bioprocess monitoring, pharmacology, toxicology, environmental sciences, bacteriology and virology. Recent advancement of flow cytometry increased its application in the reproductive biology especially for andrology. FCM is increasingly used for basic, clinical, biotechnological, and environmental studies of biochemical relevance. Although flow cytometry may overestimate the population of unlabelled cells (Petrunkina and Harrison, 2009), plethora of research from our group in pig (Pena et al., 2003, 2004, 2005; Spjuth et al., 2007; Fernando et al., 2003; Saravia et al.,2005, 2007,2009; De Ambrogi et al., 2006; ) bull (Bergquist et al., 2007; Nagy et al., 2004; Januskauskas et al., 2003; Bergqvist et al., 2007; Hallap et al., 2005, 2006;) stallion ( Kavak et al., 2003; Morrell et al., 2008) indicate that newly developed fluorescent stains and techniques of flow cytometry has made possible a more widespread analysis of semen quality at a biochemical, ultrastructural and functional level. Therefore, flow cytometry is the current technical solution for rapid, precisely reproducible assessment of sperm suspensions.
In this review we have described potentiality and scope of flow cytometry for the evaluation of semen, and the way in which this technique can be used in clinical applications for andrology based on some of our previous experiences.
Definition of flow cytometry
The definition of a flow cytometer is ‘an instrument which measures the properties of cells in a flowing stream. In other word, a flow cytometer will be defined as ‘an instrument that can measure physical, as well as multi-colour fluorescence properties of cells flowing in a stream.
In other work, cytometry refers to the measurement of physical and/or chemical characteristics of cells or, by extension, of other particles. It is a process in which such measurements are made while the cells or particles pass, preferably in single file, through the measuring apparatus in a fluid stream. The data obtained can be used to understand and monitor biological processes and develop new methods and strategies for cell detection and quantification. Compared to other analytical tools, where a single value for each parameter is obtained for the whole population, flow cytometry provides data for every particle detected. As cells differ in their metabolic or physiological states, flow cytometry allows us not only to detect a particular cell type but also to find different subpopulations according to their structural or physiological parameters.
Flow cytometry is a technique for measuring components (cells) and the properties of individual cells in liquid suspension. In essence, suspended cells are brought to a detector, one by one, by means of a flow channel. Fluidic devices under laminar flow define the trajectories and velocities that cells traverse across the detector, and fluorescence, absorbance, and light scattering are among the cell properties that can be detected. Flow sorting allows individual cells to be sorted on the basis of their measured properties, and one to three or more global properties of the cell can be measured. Flow cytometers and cell sorters make use of one or more excitation sources and one or two fluorescent dyes to measure and characterize several thousands of cells per second. Flow cytometry gives objective and accurate results (Bunthof et al., 2001; Shleeva et al., 2002), overcoming the problems with the manual methods described above.
Function and types of flow cytometry
Fluidics, optics and electronics are the three main systems that make up a flow cytometer. In a few minutes, the flow cytometer can acquire data on all subpopulations within a sample, making it ideal for assessment of heterogenous population, such as spermatozoa. The adaptation of flow cytometry to sperm assessment began when it was used for measuring their DNA content (Evenson et al., 1980) and its application to semen analysis has gradually increased over the last 10-15 years. Flow cytometry is now applied to semen evaluation of traits such as cell viability, acrosomal integrity, mitochondrial function, capacitation status, membrane fluidity and DNA status. New fluorescent stains and techniques are continuously being developed that have potential application to the flow cytometric evaluation of spermatozoa.
Flow cytometry permits the observation of physical characteristics, such as cell size, shape and internal complexity, and any component or function of the spermatozoon that can be detected by a fluorochrome or fluorescently labeled compound. The analysis is objective, has a high level of experimental repeatability and has the advantage of being able to work with small sample sizes. Flow cytometry also has the capacity to detect labeling by multiple fluorochromes associated with individual spermatozoa, meaning that more than one sperm attribute can be assessed simultaneously. This feature has an added benefit for semen analysis, as few single sperm parameters show significant correlation with fertility in vivo for semen within the acceptable range of normality (Larsson and Rodriguez-Martinez, 2000) and the more sperm parameters that can be tested, the more accurate the fertility prediction becomes (Amman and Hammerstedt, 1993).
There are two main types of flow cytometers-analysers and sorters. Sorters have the ability not only to collect data on cells (analyse cells) but also to sort cells with particular properties (defined by the flow cytometer operator) to extremely high purities. There are also a number of commercial flow cytometers that have been developed for particular analytical requirements. Partec manufacture a Ploidy Analyser and also a Cell Counter Analyser. Optoflow has developed a flow cytometer for the rapid detection, characterization and enumeration of microorganisms. Luminex is developing technology for multiplexed analyte quantitation using a combination of microspheres, flow cytometry and high speed digital processing.
Advantages of FC compared to other conventional techniques to explore sperm structure and function
During the past 2 decades, there has been an increasing interest in reliable assays for assessing semen quality in the fertility clinic and artificial insemination industries. The use of flow cytometry for sperm analysis is an attempt to address the long-standing problem of the subjective nature of the manual method commonly used for semen analysis. An additional source of laboratory variation is the low number of sperms analyzed with manual techniques. Because of time and cost restraints, most laboratories analyze only 50 to 100 sperm to compute the percentage of each cell population and the viability rate. This small sample from a population of millions probably results in a statistical sampling error (Russel and Curtis, 1993). The conventional methods used are limited to microscopic determination of sperm concentration using a hemocytometer (Jorgensen et al., 1997) and evaluation of sperm motility and morphology (Keel et al., 2002). These methods usually involve a subjective assessment of a few hundred sperm, and quality assurance is rarely implemented in the laboratories performing such analysis. Flow cytometry is a technique that is superior to conventional light microscopy techniques in terms of objectivity, number of cells measured, speed, and precision (Spano and Evenson, 1993). The technique has been used on human sperm to determine a number of factors, including membrane integrity, mitochondrial function, acrosome status, and multiparameter measurement (Garrido et al., 2002). Flow cytometry permitted us to analyze thousands of cells in few seconds. In our series of studies, we demonstrated the feasibility and reproducibility of an automated method to evaluate sperm cell type, count, and viability in human boar samples. In our hand, the precision of the flow cytometric analysis is satisfactory in diverse species (boar, bull, stallion etc), and the observed CVs were significantly better than those reported for the manual method.
While there are many advantages of using the flow cytometer for routine semen analysis, its use is often limited to research by the expense and difficulties of operation associated with the requirement of a skilled operator. In addition, a flow cytometer is quite large and cannot withstand shocks associated with movement, meaning it requires a dedicated position in the laboratory. However, the development of more affordable ‘‘bench-top'' flow cytometers has recently increased the potential application to semen analysis.
If we consider flow cytometric analysis further, we can see that it is gaining wider acceptance as a technique for assessing the acrosome reaction and viability simultaneously. Comparing these assays to the more widely used epifluorescent microscopic techniques, the flow cytometric analysis is able to give a far more simple and objective method of analysis, especially with regard to correlation of fertilization with acrosome reactivity potential (Uhler et al., 1993; Purvis et al., 1990; Carver-Ward et al., 1996).
A large number of different techniques to estimate sperm concentration have been reported. In the mid-1990s a series of fixed-depth disposable slides were evaluated as rapid and effective pieces of equipment for the estimate of sperm concentration. Preliminary data from a number of studies suggested that, at least in the 20-mm-depth format, such chambers resulted in a noticeable underestimate of sperm concentration compared to the gold standard (improved Neubauer hemocytometer). Using this information, the World Health Organization stated that ‘‘such chambers, whilst convenient in that they can be used without dilution of the specimen, may lack the accuracy and precision of the haemocytometer technique'' (World Health Organization, 1999). Further data—for example, from Tomlinson and colleagues—showed that 2 proprietary disposable slides (Microcell, Conception Technologies, San Diego, Calif; Leja, Leja Products, BV Nieuw- Vennep, The Netherlands) gave lower sperm concentrations compared to the hemocytometer method (Tomlinson et al., 2001). To put this in context, numerous reports document unacceptable discrepancies between different laboratories and even between different individuals, although fewer studies attempt to address these issues. So, what is wrong?
Several reports emphasize the need for improvement in overall quality of semen testing within and between laboratories (Neuwinger et al., 1990; Jorgensen et al., 1997; Keel et al., 2000). However, the subjective nature of conventional semen analyses, combined with their relatively low precision due to the low number of cells assessed, leads to poor intra- and interlaboratory reproducibility; therefore, the introduction of standardized or quality controlled procedures will probably have a limited effect. The conventional analyses are used to determine whether parameters obtained from an ejaculate are within the range characterized by fertile men, and these methods can therefore provide only unclear cut-off values when used for the prediction of fertility status. Many of the advantages that accrue when using flow cytometry may, when applied to assessment of sperm cells, help overcome some of the mentioned problems found in conventional semen analysis.
In the field of semen analysis, validation of a method is important because it is essential to have specific, precise, objective, and accurate laboratory tests to establish a correlation of the data with fertility or to determine the fertility potential of a semen sample correctly (Amann, 1989). Precision of a laboratory test is of great concern to the andrologist in the fertility clinic, since the results of the semen analysis are often used to advise a patient about his fertility and the prognosis for the treatment of the couple. To use established cut-off values and ensure uniform diagnosis, within and between laboratory variations should be determined and followed closely.
Accurate determination of sperm cell concentration is critical to the AI industry because it provides assurance both to bull studs and to customers that straws of extended semen contain the sperm numbers indicated. An accurate measure of sperm concentration is particularly important in export markets in which verification of numbers may be required. Routine sperm counts can help to identify possible processing errors within a specific batch of semen or on a particular day, should those errors occur. As sperm counting procedures become more refined, routine counting can be used to monitor subtle changes in daily semen processing that might affect the number of sperm packaged in a straw.
Hemacytometers are widely used for routine sperm counts, but the equipment is slow, and multiple measurements of each sample are needed. Single hemacytometer counts are not highly accurate; because of inherent errors in the technique, Freund and Carol (13) found that mean differences of 20% were not uncommon between duplicate sperm count determinations by the same technician. Electronic counters provide much more rapid counting, are easier to use, and give more repeatable results among technicians. However, those instruments tend to include in the sperm count any somatic cells present, immature sperm forms, cytoplasmic droplets, debris, and bacteria, thereby inflating the concentration value (19).
Currently, the primary method used by the AI industry to estimate sperm concentration is spectrophotometric determination of turbidity of a semen sample using an instrument previously calibrated for sperm concentration with a hemacytometer or Coulter counter (1). This approach is only as accurate as the methods used for spectrophotometer calibration.
New, more accurate methods for sperm count determinations are being sought to replace the older ones. Some laboratories are trying the Maklerm counting chamber (Seif- Medical, Haifa, Israel) and other improved hemacytometers, such as the MicroCellTM (Fertility Technologies, Inc., Natick, MA); however, these techniques will likely have standard lems similar to those associated with the standard hemacytometers.
It may be argued that when comparing fluorescent microscopy assays with flow cytometry, one is examining "patterns" of fluorescence rather than fluore'scence intensity, i.e., the flow cytometer is not capable of discriminating sperm which have a fluorescent marker bound to the equatorial segment or over one of the acrosomal membranes (Parinaud et al., 1993; Mortimer and Camenzind, 1989; Mortimer et al., 1987). Tao et al. (1993) compared flow cytometry and epifluorescent microscopy with various lectins and indicated that there is no significant difference between the two methodologies for detection of the acrosome reaction. However, it has been argued that lectins do not bind specifically to the acrosomal region of the sperm (Purvis et al., 1990; Holden and Trounson, 1991) and that other binding sites can be easily distinguished by epifluorescence microscopy, whereas flow cytometry identifies the signal from the entire sperm.
Additionally, conventional light microscopic semen assessment is increasingly being replaced by fluorescent staining techniques, computer-assisted sperm analysis (CASA) systems, and flow cytometry (Pen˜a et al., 2001; Verstegen et al., 2002). Additional advantages over existing techniques are that this approach is faster than the hemacytometer and that cellular debris, fat droplets, and other particulate material in extended semen are not erroneously counted as sperm, as often occurs with electronic cell counters. This method can also be used to determine the number of somatic cells in a semen sample.
The viability of spermatozoa is a key determinant of sperm quality and prerequisite for successful fertilization. Viability of spermatozoa can be assessed by numerous methods, but many are slow and poorly repeatable and subjectively assess only 100 to 200 spermatozoa per ejaculate. Merkies et al. (2000) compared different methods of viability evaluation. They concluded that Eosin-nigrosin overestimate viability while fluorescent microscope and flow cytometry estimate similar trend of viability. Currently flow cytometric procedures have been developed which simultaneously evaluate sperm cell viability, acrosomal integrity and mitochondrial function. This method has been successfully used for assessing spermatozoa viability in men (Garner and Johnson, 1995), bulls (Garner et al., 1994; Thomas et al., 1998), boars (Rodríguez-Martínez, 2007; Garner and Johnson, 1995; Garner et al., 1996), rams (Garner and Johnson, 1995), rabbits (Garner and Johnson, 1995), mice (Garner and Johnson, 1995; Songsasen et al., 1997), poultry and wildfowl (Donoghue et al., 1995; Blanco et al., 2000) and honey bees (Collins and Donoghue, 1999; Collins, 2000) and in fish (Martin Flajshans et al., 2004).
Considerable information has accumulated on the use of fluorescent staining protocols for assessing sperm viability (Evenson et al., 1982). The SYBR 14 staining of nucleic acids, especially in the sperm head, was very bright in living sperm. Good agreement was observed between the fluorescent staining method and the standard eosin-nigrosine viability test; the flow cytometric method showed a precision level higher than that of the manual method.
One of the first attempts to assess sperm viability utilized rhodamine 123 (R123) to assess mitochondrial membrane potential and ethidium bromide to determine membrane integrity using flow cytometry (Garner et al., 1986). Other combinations that have been used to examine the functional capacity of sperm are carboxyfluorescein diacetate (CFDA) and propidium iodide (PI) (Garner et al., 1988; Watson et al., 1992); carboxydimethylfluorescein diacetate (CMFDA), R123, and PI (Ericsson et al., 1993; Thomas and Garner, 1994); and PI, pisum sativum agglutinin (PSA), and R123 (Graham et al., 1990).
At present, one of the most commonly used viability stain combinations is SYBR-14 and PI, sold commercially as LIVE/DEAD Sperm Viability kit (Molecular Probes Inc., OR, USA). When used in combination, the nuclei of living sperm fluoresce green (SYBR-14) and cells that have lost their membrane integrity stain red (PI). This staining technique has been used in a number of species, including the boar (Garner and Johnson, 1995; Saravia et al.,2005, 2007,2009). Although species differences do exist in the function of spermatozoa, the Live/Dead stain may similarly have no adverse affect on fertilization in the equine, although it remains to be tested in this species. Recently a new instrument (Nucelocounter-SP100) has been used to evaluate boar sperm concentration . Due to its compact size and its relatively inexpensive purchase price, this instrument could be useful for field measurements of both concentration and viability. This instrument was considered to be a useful instrument for rapidly measuring stallion sperm concentration and viability (Morrell et al., 2010).
Fluorescent probes such as H33258, requiring flow cytometric analysis with a laser that operates in the ultraviolet light range, are less commonly used as this is not a standard feature on the smaller analytical machines. However, one alternative is to use a fluorometer. A fluorometer is a relatively low-cost piece of portable equipment that permits a rapid analysis to be carried out on a sample. Januskauskas et al. (2001) used H33258 to detect nonviable bull spermatozoa by fluorometry and found a negative correlation between the percentage of damaged cells and field fertility. Another option is fluorescent attachments for computer-assisted semen analysis devices. For example, the IDENT fluorescence feature of the Hamilton-Thorne IVOS permits staining with H33258 allowing an assessment of sperm viability to be made along with motility.
Fluorochromes used to assess sperm viability by either approach can be used in combination with each other. For example, when CFDA is used along with PI, three populations of cells can be identified: live, which are green; dead, which are red; and a third population which is stained with both and represents dying spermatozoa. Almlid and Johnson (1988) found this combination useful for monitoring membrane damage in frozen-thawed boar spermatozoa during evaluation of various freezing protocols. Harrison and Vickers (1990) also used this combination with a fluorescent microscope and found it to be an effective indicator of the viability of fresh, incubated or cold-shocked boar and ram spermatozoa. Garner et al. (1986) used this combination to stain spermatozoa from a number of species, but at that time could not find a relationship between bull sperm viability detected by CFDA/PI and fertility.
Flow cytometry for assessment of sperm viability appears to be a valuable tool for the AI industry. When a high number of sperm is packed in each insemination dose, the effect of selecting the best ejaculates according to sperm viability has a relatively limited effect on NRR56. However, sperm viability might be more important when combined with low-dose inseminations. The FACSCount AF flow cytometer also determines sperm concentration accurately and precisely during the same analysis (Christensen et al., 2004a). The combination of assessment of sperm viability and concentration appears to be useful in the improvement of quality control at AI stations. Because of the results of this trial, this method has been implemented by Danish AI stations (Christensen et al., 2005). Relatively bright fluorescence was found also in the mitochondrial sheath of living sperm. The mechanism by which SYBR-14 binds to the DNA is not known. It is know that PI stains nucleic acids by intercalating between the base pairs (Krishan, 1975). Viability stains have also been used in association with fluorescently labeled plant lectins to simultaneously assess the plasma membrane integrity and the acrosome integrity (Nagy et al., 2003). Assessment of viability using SYBR-14 dye does not damage spermatozoa, since Garner et al. (5) demonstrated that insemination of boar spermatozoa stained with SYBR-14 into sows did not compromise fertilization or the development of flushed porcine embryos in culture.
Non-viable cells can be determined using membrane-impermeable nucleic acid stains which positively identify dead spermatozoa by penetrating cells with damaged membranes. An intact plasma membrane will prevent these products from entering the spermatozoa and staining the nucleus. Commonly used examples include phenanthridines, for example propidium iodide (PI; (Matyus, 1984) ethidium homodimer-1 (EthD-1; (Althouse et al., 1995), the cyanine Yo-Pro (Kavak, 2003) and the bizbenzimidazole Hoechst 33258 (Gundersen and Shapiro, 1984). Wilhelm et al. (1996) compared the fertility of cryopreserved stallion spermatozoa with a number of laboratory assessments of semen quality and found that viability, as assessed by flow cytometry using PI, was the single laboratory assay that correlated with stallion fertility.
Changes in sperm membrane permeability
Detection of increased membrane permeability is employed in different cell types to distinguish different status of membrane organization (Cohen, 1993; Ormerod et al., 1993; Castaneda and Kinne, 2000; Reber et al., 2002). Sperm plasma membrane status is of utmost importance due to its role, not only as a cell boundary, but also for its need for cell-to-cell interactions, e.g. between spermatozoa and the epithelium of the female genital tract and between the spermatozoon and the oocyte and its vestments (for review, see Rodriguez-Martinez, 2001). Membrane integrity and the stability of its semipermeable features are prerequisites for the viability of the spermatozoon (Rodriguez-Martinez, 2006). However, cryopreservation, whose purpose is to warrant sperm survival, causes irreversible damage to the plasma membrane leading to cell death in a large number of spermatozoa (Holt, 2000) or, in the surviving spermatozoa, to changes similar to those seen during sperm capacitation, thus shortening their lifetime (Perez et al., 1996; Cormier et al., 1997; Maxwell and Johnson, 1997; Green and Watson, 2000; Schembri et al., 2000; Watson, 2000). During the freezing process, cells shrink again when cooling rates are slow enough to prevent intracellular ice formation as growing extracellular ice concentrates the solutes in the diminishing volume of non-frozen water, causing intracellular water exosmosis. Though warming and thawing, the cells return to their normal volume. Thus, it is important to know the permeability coefficient of the cells to cryoprotectants, as well as the effect of cryoprotective agents on the membrane hydraulic conductivity.
Classical combination of probes allows discrimination of two or three subpopulations of spermatozoa, i.e. live, dead and damaged depending on the degree of membrane integrity (Eriksson & RodrÄ±´guez-MartÄ±´nez, 2000). A new, simple and repeatable method to detect membrane changes in all spermatozoa present in a boar semen sample, by use of markers (combination of SNARF-1, YO-PRO-1 and ethidium homodimer) used to track changes in sperm membrane permeability, has been developed recently by our group (Pena et al., 2005). In determined physiological or pathological situations, live cells are unable to exclude YO-PRO-1, but are still not permeable to other dead-cell discriminatory dyes, like propidium iodide or ethidium homodimer. YO-PRO-1 is an impermeable membrane probe and can leak in, only after destabilization of the membrane, under conditions where ethidium homodimer does not. Because several ATP-dependent channels have been detected in spermatozoa (Acevedo et al., 2006), it seems plausible that this is a result of the silencing of a multidrug transporter. This multidrug transporter is involved in transporting amphipathic small molecules like YO-PRO-1, which in intact cells is actively pumped out but not after destabilization of the plasma membrane, maybe because sub-viable cells lack appropriate amounts of ATP to transport YO-PRO-1 back out of the cell (Ormerod et al., 1993). Therefore, the use of a fluorescent probe, such as YO-PRO-1, which penetrates cells as they undergo changes related to cryoinjury, where membranes become slightly permeable, makes YO-PRO-1 a useful tool for detecting early membrane changes (Idziorek et al., 1995; Wronski et al., 2000). This triple staining distinguishes, as in the Annexin assay, four sperm subpopulations. The three probes are easily distinguished both in flow cytometry and in fluorescence microscopy. The absorption and emission maxima for YO-PRO-1 are 491 nm and 509 nm, respectively, and 528 nm and 617 nm, respectively, for ethidium homodimer to be detected in the Flow Cytometer with the FL1 and FL3 photomul-tipliers.
The triple staining technique offers some advantages over the A/PI assay. Whereas in the A/PI assay there is always an unstained subpopulation, the triple stain labels all the spermatozoa in the sample, an obvious advantage when using manual counting in fluorescence microscopy. If a flow cytometer is available, because only sperm cells are stained with the triple staining technique, spermatozoa and debris can be easily separated based not only on scatter properties of the particles but also on their fluorescent properties. This fact is important because in bull semen, it has been demonstrated that egg-yolk particles can be easily misjudged as spermatozoa based only on their scatter properties (Nagy et al., 2003), requiring preliminary washing and centrifugation to cleanse the cells. Centrifugation might cause sperm damage and, therefore, mask other effects caused by the cryopreservation. The subpopulation of live cells using the new triple staining concurs with the subpopulation of live cells using the well validated A/PI assay. In addition, the staining protocol is much easier than the A/PI because the staining is made from stock solutions and is not necessary to use a binding buffer. As the staining of the probes is not dependent on Ca2+, as is the case binding FITC-A, the preparation and using of a Ca2+ enriched buffer is not necessary. The agreement between both techniques (A and YOPRO- 1/Eth/SNARF-1) was good, although the percentage of live spermatozoa was slightly higher in the triple staining method (Pena et al., 2006). Also, the percentage of early damaged spermatozoa was higher with the A/PI assay. This might reflect an increase in membrane permeability, preceeding the transposition of PS in the evolution of the cryodamage, or in a yet to be determined physiological change probably being a very early step of both processes related to changes in cell volume regulation and movement of ions, occurring during the initiation of apoptosis (Bortner and Cidlowski, 1998) or cryoinjury (Paasch et al., 2005). In addition, an earlier inactivation of enzymes involved in maintaining membrane asymmetry than those involved in transporting amphipatic small molecules like YO-PRO-1 might explain this fact.
Sperm plasma membrane integrity
Although the sperm plasma membrane covers the entire cell, it consists of three distinct membrane compartments, one which covers the outer acrosomal membrane, one which covers the post acrosomal portion of the sperm head, and one which covers the middle and principal pieces. The numerous functions of the membrane are related to the cell metabolism for maintaining sperm motility, capacitation, acrosome reaction, interactions between the spermatozoa and the epithelium of the female genital tract, and sperm-egg interactions (Rodriguez-Martinez, 2003). Most ‘viability assays' assess whether or not the plasma membrane is intact (the cell is ‘viable') or not (the cell is ‘dead'). However, because the plasma membrane is composed of these different compartments, different viability assays assess the integrity of different plasma membrane compartments. Classical stains, such as eosin-nigrosin and eosin aniline blue, as well as more recent fluorescent stains, such as propidium iodide, ethidium bromide, 4-6-diamidino-2-phenylindole (DAPI) and bisbenzimide (Hoechst dyes), bind to and stain the DNA of sperm that possess a post acrosomal plasma membrane that is not intact. However, these probes will not assess the integrity of the plasma membrane covering the acrosome or principal piece. The integrity of the plasma membrane covering the principal piece can be assessed using sperm motility or the hypo-osmotic swelling test (Jeyendran et al., 1984; Neild et al., 2000; Colenbrander et al., 2003). The integrity of the plasma membrane covering the acrosome is generally assessed in conjunction with the integrity of the outer acrosomal membrane. The acrosomal ridge, present on the sperm of several species, can be used to assess the integrity of this plasma membrane by differential interference phase contrast microscopy, but this anatomical component can only be used for sperm from those species which possess it. Several non-fluorescent and fluorescent staining combinations (reviewed by Cross and Meizel, 1989; Graham, 2001; Colenbrander et al., 2003; Silva and Gadella, 2006) have been developed to permit assessment of acrosomal membrane integrity of fresh and fixed sperm samples using microscopy, fluorometry and flow cytometry.
During the last decade, several fluorescent dyes were used and validated for the assessment of the sperm membrane integrity in dogs: carboxyfluorescein diacetate (CFDA) in combination with propidium iodide (PI) (Pen˜a et al., Rota et al., 1995), SYBR-14 in combination with PI (Rijsselaere et al., 2002; Yu et al., 2002), carboxy-seminaphthorhodfluor (Carboxy-SNARF) in combination with PI (Pen˜a et al., 1999), calcein-AM in combination with ethidium homodimer (Calcein-AM/EthD-1) (Sirivaidyapong et al., 2000) and Hoechst 33258 (Hewitt and England, 1998). The major advantage of fluorescent staining techniques is the possibility to analyse fluorescently labelled spermatozoa by means of flow cytometry, enabling the evaluation of larger numbers of spermatozoa in a short interval. Pen˜a et al. (1998, 2001) found high correlations between flow cytometry and epifluorescence microscopy for the percentage of live and dead spermatozoa as determined by a CFDA-PI staining.
Apoptosis is a carefully regulated process of cell death that occurs as a normal component of development. During apoptosis, the plasma membrane became slightly permeable and loses asymmetry in one of the earliest stages of the process. When the cell membrane is disturbed the phospholipid PS is translocated from the inner to the outer leaflet of the plasma membrane (Desagher and Martinou, 2000). Recently, it has demonstrated that freezing-thawing of human (Kemal et al., 2001) and bull (Anzar et al., 2002) semen induces membrane PS translocation, thus indicating cryopreservation is a cause of apoptosis (Baust, 2002). During the early phases of disturbed membrane function, asymmetry of the membrane phospholipids occurs, before the integrity of the plasma membrane is progressively damaged (11). In all mammalian cell types studied, including spermatozoa, the two leaflets of the plasma membrane bilayer differ in phospholipid content. Phosphatidylserine (PS) and phosphatidylethanolamine (PE) are concentrated in the inner leaflet, whereas sphyngomyeline (SM) and phosphatidylcholine (PC) are concentrated in the outer leaflet (12).
The evaluation of sperm membranes is an appropriate indicator of the success of cryopreservation since sperm membrane are extremely susceptible to cryoinjury (4-6). Deleterious effects of boar sperm manipulation such as excessive extension, sorting, chilling or cryopreservation that lead to membrane destabilization. Freezing and thawing procedures significantly affect the lipid composition of boar sperm plasma membranes (34), even in the presence of good sperm quality. The negative effects of cooling, freezing and thawing are mainly caused by lipid phase transitions, ice crystallization and osmotic-induced water fluxes, and subsequent membrane recognizations influence membrane integrity, structure and function (Hammerstedt et al., 1990).
Compared with somatic cells, sperm plasma membranes have especially high levels of long-chain polyunsaturated fatty acids. Recently, variations in PUFAs within sperm plasma membranes, in particular DPA and DHA, have been associated with differences in cryotolerance in sperm isolated from Asian and African elephants (Swain & Miller 2000), common wombats, grey kangaroos and koalas (Miller et al., 2004), and blue foxes and silver foxes (Miller et al., 2005).
Differences in cryotolerance have previously been related to a high cholesterol/phospholipid ratio and low unsaturated/saturated fatty acids ratio in sperm membranes; however this relationship cannot explain the observed differences in cryotolerance between species such as bull and boar, or between individuals. The terms ‘good freezer' and ‘bad freezer' have existed for a long time, and it seems to be a trait of the individual animal rather than a characteristic of individual ejaculates. In fact, newly published results from Thurston et al. (2002) indicate that there is a genetic basis for the variation in cryopreservation-induced injuries, which included disrupted plasma and acrosomal membranes and poor motility, between individuals categorized as ‘good freezers' and those categorized as ‘bad freezers'. While the underlying mechanism(s) for the genetic differences related to cryopreservation- induced injuries is unknown, it has been suggested that these male-to-male differences may represent differences in sperm lipid and protein composition (Holt et al., 2005). These include addition of cryoprotectants prior to freezing, volumetric changes and associated membrane stretching and shrinkage in response to hyperosmotic cryoprotectant solutions as well as freeze-induced dehydration, thermotropic and lyotropic phase transitions in membrane phospholipids, and the well-established effects of elevated solute concentration and intracellular ice formation which are cooling rate-dependent.
The acrosome is a membrane enclosed structure covering the anterior part of the sperm nucleus. Powerful hydrolyzing enzymes belongs to that structure, a basic feature of the sperm head of all mammals (Yanagimachi, 1994). The acrosome reaction (AR) of mammalian sperm is to be observed before fertilization, as it is necessary for sperm penetration of the zona pellucid and for fusion with plasma membrane. As a prerequisite of fertilization the content of the acrosome is released into its surroundings during the acrosome reaction. It is assumed that the AR serves at least dual functions by facilitating the ability of the sperm to penetrate the ZP, and subsequently by aiding in the oocyte-sperm fusion process (Yanagimachi, 1994). Acrosomal loss can also occur in degenerating (dying) spermatozoa, it is considered essential to distinguish between the occurrence of acrosome reaction in viable and non viable cells (Cross and Meizel, 1989). To take account of this in multiparametric studies, various dyeing methods have been developed to assess AR. Hence, the efficacy of a sperm population to undergo the AR could be expected to influence male fertilizing potential.
Acrosome intactness, a prerequisite for fertilisation, can be readily examined in vitro using phase-contrast microscopy (Rodríguez-Martínez et al. 1997a). Most frequently used methods are triple or double staining (Talbot and Chacon, 1980; De Jonge et al., 1989), isothiocyano-fluoresceinated Pisum sativum agglutinin (FITC-PSA; Crosset al., 1986), FITC-concanavalin A (FITC-ConA; Holden et al., 1990), Arachis hypogaea agglutinin (FITC-peanut agglutinin FITC-PNA; Kallajoki et al., 1986; Mortimer et al., 1987), chlortetracycline (Saling and Storey, 1979; Amin et al., 1996), paramagnetic beads (Okabe et al., 1992; Ohashi et al., 1992, 1994), Coomassie Blue (Aarons et al., 1993), anti-acrosin antiserum (Tesarik et al., 1990), mannosylated bovine serum albumin (Benoff et al., 1993), quinacrine (Amin et al., 1996) and monoclonal antibodies (Kallajoki and Suominen, 1984; Wolf et al., 1985; Moore et al., 1987; Fe´nichel et al., 1989; Aitken and Brindle, 1993; Chao et al., 1993). A combined assessment method for human sperm morphology and the acrosomal status was demonstrated by O'Bryan et al. (1994), who used monoclonal antibodies against clusterin. This glyco-protein is located within the acrosomal cap. Electron microscopic studies have been performed to compare the ultrastructural morphology of acrosomes with the staining patterns of spermatozoa after labelling with the antibodies by the Köhn et al. (1997). There are some limitations of microscopic examination it is only easily accomplished in species such as the hamster and guinea pig which have large acrosomes. However, most mammalian sperm, including those of humans have such small acrosomes thus normal reactions are not easily observed with the light microscopy. Although epifluorescence microscopy is being used, Pen˜a et al.  found that epifluorescence microscopy was less precise than flow cytometry for detecting the percentage of spermatozoa with damaged acrosomes, probably due to the difference in sample size. Nevertheless, Miyazaki et al. (1990) stated that percentage of acrosome-reacted sperm determined by flow cytometry and fluorescence microscopy showed that these methods gave very similar results.
The advent of cellular measurements by flow cytometric analysis of individual attributes within a population of cells is an important step toward the evaluation of acrosome reaction. Comparing other assays to the more widely used epifluorescent microscopic techniques, the flow cytometric analysis is able to give a far more simple, speedy, accurate and objective method of analysis, especially with regard to correlation of fertilization with acrosome reactivity potential (Uhler et al., 1993; Purvis et al., 1990; Carver-Ward, 1996). High levels of green and red fluorescence are characteristic of non-reacted spermatozoa, while the AR produce a decreased fluorescence intensity. FC analysis has permitted determination of the regions from reacted and non-reacted populations, and thus calculation of the percentage for each. This determination is normally corroborated by fluorescence microscopy observations. Acrosomal integrity can be measured by a number of methods, but the most commonly used method is with a plant lectin labeled by a fluorescent probe. There are a large number of lectins available for assessing acrosomal integrity, some of which are toxic such as Ricinus communis agglutinin. Pisum sativum agglutinin (PSA) is a lectin from the pea plant that binds to a-mannose and agalactose moieties of the acrosomal matrix. Since PSA cannot penetrate an intact acrosomal membrane, only acrosome-reacted or damaged spermatozoa will stain (Cross et al., 1986). Arachis hypogaea agglutinin (PNA) is a lectin from the peanut plant that binds to bgalactose moieties associated with the outer acrosomal membrane of fixed spermatozoa, indicating acrosome-intact cells (Mortimer et al., 1987). A hypogaea agglutinin is believed to display less non-specific binding to other areas of the spermatozoon, leading some workers to favour this over PSA (Graham, 2001). Carver-Ward et al. (1997) proposed that PNA is the reliable lectin compared to PSA, CD46 and ConA. Thus, PNA is capable of differentiating the acrosome reacted sperm from a given population. However, in observing that only PNA gives a specific comparison between non-acrosome- reacted and acrosome-reacted sperm, the differences between the two markers are merely a matter of magnitude. Tao et al. (36) also state that PNA is a more reliable acrosome reaction marker compared to PSA, Con A, and SBA. Petrunkina et al. (2005) observed that PNA-FITC, which was used in this study to evaluate sperm responsiveness, binds to the outer acrosomal membrane (OAM) (Fazeli et al., 1997; Flesch et al., 1998), so that FITC-PNA can be used as a probe to monitor boar sperm acrosome reaction.
One of the most used fluorochrome combinations for simultaneous evaluation of plasma membrane integrity (i.e., viability) and acrosome integrity are fluorescein isothiocyanate- conjugated pea (Pisum sativum) agglutinin (FITC-PSA) and propidium iodide (PI) (Graham et al., 1990). On nonpermeabilized spermatozoa, FITC-PSA provides information regarding the integrity of the acrosome. Sperm cells with an intact acrosome will have no fluorescence, and cells with a reacted or damaged acrosome will show green fluorescence. Propidium iodide is a DNA-specific stain that cannot enter the intact plasma membrane and, therefore, is used as a dead-marker counterstain. This double-staining for sperm viability and acrosome integrity is relatively reliable for fresh and in vitro-capacitated sperm, because sperm cell particles can easily be distinguished from nonsperm events by their specific forward- and sideways-scatter properties e.g., in dogs (Szasz et al., 2000). For frozen-thawed sperm, the main problem is that many of the egg yolk particles have scatter properties similar to those of sperm cells that trouble the elimination of nonsperm events by scatter gating enormously (Pena et al., 1999). Such egg yolk particles, like live acrosome-intact sperm cells, have low fluorescence and, therefore, will be assessed as live acrosome-intact sperm using the PI/FITCPSA double-labeling method. Therefore, when using the PI/FITC-PSA double-staining protocol, complete removal of yolk particles from thawed sperm suspensions is required for accurate analyses of sperm integrity after cryopreservation. However, it has been argued that lectins do not bind specifically to the acrosomal region of the sperm (Purvis et al., 1990; Holden and Trounson, 1991) and that other binding sites can be easily distinguished by epifluorescence microscopy, whereas flow cytometry identifies the signal from the entire sperm. From a biological point of view, both the plasma membrane and the outer acrosomal membranes fuse and vesiculate during the acrosome reaction, which would preclude binding to the outer acrosomal membrane on nonpermeabilized sperm. Also, it is questionable as to how one could permeabilize the plasma membrane without affecting the underlying outer acrosomal membrane. AR in spermatozoa can also be detected with FC using monoclonal antibodies against inner acrosomal membrane epitopes such as GB24, MH61 and CD46 (48-53). Tao et al. (1993) examined the use of two monoclonal antibodies (MH61 and CD46) for acrosome reaction assessment, while others have concentrated on CD46 alone (D'Cruz and Haas, 1992; Carver-Ward et al., 1994).
Maxwell and Johnson (1997) treated boar spermatozoa with lysophosphatidylcholine in order to mimic the accumulation of lysophospholipid, which has been implicated in the acrosome reaction, and found a significant increase in FITC-PSA spermatozoa when assessed by flow cytometry. Pena et al. (2001) stated that compared to microscopic evaluation, the results demonstrated that flow cytometry is a precise method for evaluating the viability and acrosomal status of fresh samples of dog semen. A new triple staining procedure, using carboxy-SNARF-1, propidium iodide and FITC-PSA, was developed and was an efficient method for evaluating actosomal integrity of cryopreservation protocols for dog spermatozoa. Herrera et al. (2002) used FITC-PSA to determine whether there was an association between the acrosome reaction and the incidence of subfertility of boar spermatozoa. The authors found that the percentage of spontaneous acrosome reaction was not significantly different in fertile and subfertile boars. However, the incidence of a progesterone-induced acrosome reaction was significantly lower in subfertile (5.75%) compared with fertile boars (10.0%), suggesting that assessment of the induced acrosome reaction may be a useful parameter to assess fertility. In a similar study, Jime´nez et al. (2002) attempted to determine the kinetics of surface carbohydrate turnover during in vitro capacitation and the acrosome reaction in fertile and subfertile boars. Spermatozoa were exposed to three FITC-labeled lectins: Triticum vulgaris agglutinin (WGA; specific for sialic acid and N-acetylglucosaminyl residues), Concanavalia ensiformis agglutinin (Con- A; specific for D-mannosyl and D-glucosyl residues) and Ulex europaeus agglutinin (UEA; specific for L-fucose), and assessed by flow cytometry. The authors reported differences in lectin patterns across capacitated and acrosome-reacted spermatozoa between fertile and subfertile boars.
From a study of Cooper and Yeung (1998) use of F-fucoidin as vital dye for flow cytometry analysis was emphasized. They suggested that F-fucoidin binds not to the plasma membrane of intact cells but rather to intraacrosomal components, presumably exposed in cells with damaged membranes or those that have undergone the acrosome reaction.
Capacitation of sperm is a prerequisite for successful fertilization. Capacitation is an important, but rather incompletely understood phenomenon that a spermatozoon undergoes before it can fertilize the oocyte. Capacitation is reversible and lasts hours. During this process various cellular changes occur at specific times and locations, including an increase in membrane fluidity due to lipid modifications, an influx of calcium to the sperm head and flagellum, the generation of controlled amounts of reactive oxygen species, as well as the phosphorylation of proteins on serine, threonine and tyrosine residues (De Jonge, 2005; O'Flaherty et al., 2006; Lamirande and O'Flaherty, 2007; Tulsiani et al., 2007). Therefore, we are now able to evaluate the ability of spermatozoa to capacitate under various conditions, which should provide information regarding cell longevity and performance. Capacitation is associated with alterations in a variety of intracellular and sperm surface features, but the precise relationship between these modifications and capacitation is not certain. Therefore, in spite of many published reports describing phenomena that might be correlated with capacitation, there is no general consent as to the assay (s) or techniques that can distinguish capacitated from noncapacitated spermatozoa.
One of the most often used methods for determination of the capacitation status is the CTC (chlortetracycline) assay by using fluorescence microscopy. This fluorescent antibiotic will detect and exhibits enhanced fluorescence over the segments of the membrane where Ca2+ accumulates. Chlortetracycline has been shown to interact with mammalian spermatozoa, showing different binding patterns on the sperm head, which are believed to reflect different stages of the capacitation process (Ward&Storey, 1984; DasGupta et al., 1993; Fraser et al., 1995). Though, CTC is empirically accepted but is laborious to use and its working mechanism is scientifically unexplained.
Although sperm capacitation is not only a destabilization process, early stages of sperm capacitation can be measured by loading spermatozoa with the lipid dye merocyanine-540 (Harrison and Gadella, 2005) and then using flow cytometry to determine any significant increase in fluorescence (related to the degree of lipid disorder in the plasma membrane and indicative of of the beginning of capacitation; Redriguez-Martinez et al., 2001). Flow cytometric detection of capacitation-related changes in membrane architecture using merocyanine 540 as a reporter probe, and acrosome status using FITC-PNA staining, have some clear advantages over the all-compassing CTC staining technique. First, given the clear differences in the intensity of fluorescence between control and capacitated or acrosome-reacted cells, flow cytometry allows for the very rapid and objective discrimination of the status of large numbers of sperm cells. For example, in the current study we analyzed 10 000 sperm cells per data point in only a few seconds. Second, prior to analysis, the sperm suspension requires only simultaneous addition of appropriate amounts of PI and FITC-PNA or Yo-Pro-1 and merocyanine 540, followed by a 10-min incubation for the completion of labeling. Third, the cells can be analyzed in a flow cytometer in the unfixed state and under relatively physiological conditions. This ability to control the ambient conditions minimizes the risk of cell deterioration, especially for the notoriously delicate capacitated sperm cells. Several recent studies have found a correlation between sperm capacitation and phosphorylation on tyrosine residues of specific sperm proteins suggesting that measurement of protein tyrosine phosphorylation may give a quantitative estimate of capacitation. Sidhu et al. (2004) has described a new method for estimating levels of tyrosine phosphorylation in spermatozoa undergoing capacitation. The global levels of sperm tyrosine phosphorylation under different in vitro conditions that induce capacitation were estimated in permeabilized cells using FITC-conjugated mAbs against tyrosine phosphoproteins and flow cytometric analysis. This technique is rapid, simple and reliable. The specificity of the technique was demonstrated by specific displacement of FITC-labeled PT66 mAb by phosphotyrosine and the failure of PT66 to label non-permeabilized intact sperm owing to the intracellular localization of phosphoproteins. Staurosporine, a strong inhibitor of the phosphotyrosine kinase enzyme specifically inhibited sperm tyrosine phosphorylation.
Mitochondria, located in the sperm midpiece, are the primary energy generating centers for motility and other processes in the sperm cell as in most other cells. Interference with this function has severe consequences for any cell (Krahenbuhl, 2001). A number of crucial events in apoptosis commence in the mitochondria (Zamzami et al., 1995; Lui et al., 1996; Petit et al., 1996; Li et al., 1997). Mitochondria ensheath the midpiece of the spermatozoa and deliver adenosine triphosphate (ATP) to the axenome where it is utilized for flagellar propulsion. These organelles are required for efficient energy metabolism, production of membrane lipids and cell growth but are also the primary determinants of cellular life or death (Arends and Wyllie, 1991). In other words, mitochondria have been verified as the co-ordinators of apoptosis in numerous cell systems (Frade and Michaelidis, 1997; Kroemer, 1997; Zhyng et al., 1998). Shivaji et al. (2009) focuses on the identity and function of mitochondrial proteins which undergo capacitation-dependent tyrosine phosphorylation in spermatozoa. Hallap et al. (2005) reported that the results obtained from flow cytometric measurements of mitochondrial function were 10-15% lower than the recordings of motility, either subjectively or as measured by CASA. Such a difference between subjective motility evaluations and flow cytometry evaluations of mitochondrial activity is in line with that reported in several other publications (Garner et al., 1997; Gravance et al., 2000; Wu et al., 2003).
Accumulation in mitochondria is characteristic of many fluorescent dyes, such as Rhodamine 123 (R123), MitoTracker Green (MTG), JC-1, MitoTracker Orange (CMTMRos), MitoTracker Red (CMXRos), MitoTracker Red 580, and MitoTracker Deep Red 633 (Cossarizza---; Garner et al., 1997; Gravance et al., 2002; Ericsson et al., 1993). Despite of numerous published protocols, there are several problems connected with most of these fluorophores. There are approximately 100 mitochondria in the mid-piece of the spermatozoon and fluorescent dyes, able to target defined intracellular compartments, can be used to visualize them. Most of these dyes work by diffusing into living cells and accumulating in mitochondria, provided that an internal 100-200 mV negative potential gradient occurs across the mitochondrial membrane (MMP).
The most widely used mitochondrial-specific probe, R123 is a cationic compound that excites at 488 nm and emits at 515-575 nm (green fluorescence). It accumulates in the mitochondria as a function of transmembrane potential (Chen, 1988; Al-Rubeai, 1993), of R123 concentration, and of sperm numbers (Windsor and White, 1993); it is not dependent on time or temperature (Auger et al., 1989). It was historically applied to spermatozoa in combination with ethidium bromide (Evenson et al., 1982). The R123 accumulated in the mitochondria and fluoresced green, thus identifying the sperm that exhibited a mitochondrial membrane potential. The dead spermatozoa, such as those with damaged membranes, were identified by the uptake of ethidium bromide. A similar combination, R123 and propidium iodide (PI), has been shown to readily discriminate between living and dead spermatozoa (Evenson et al., 1982). Although functioning mitochondria stain green with rhodamine123, this stain does not permit one to differentiate between mitochondria that exhibit different respiratory rates. The novel mitochondrial probe, MitoTracker Green FM (MITO), is nonfluorescent in aqueous solution and fluoresces green upon accumulation in the mitochondria regardless of mitochondrial membrane potential (Haugland, 1996). R123 and MITO are transported into actively respiring mitochondria and their accumulation in the mitochondria causes them to fluoresce green. R123 is not suitable for use in experiments in which the spermatozoa are treated with aldehyde fixatives, whereas the MITO probes are well retained during the fixation process. MITO-labeled mouse spermatozoa were placed in the female reproductive tract by AI to trace the distribution of the mitochondria in the developing embryo (Davies and Gardner, 2002). Gadella and Harrison (2002) also used MITO to show that bicarbonate does not affect the mitochondrial potential of boar spermatozoa. Fluorescence-activated flow cytometric assays of Jagg fluorescence have been used to identify sperm with high MMP in a number of experiments with sperm (Garner et al., 1997; Gravance et al., 2001; Love et al., 2003). Results of Guthrie and Welch (2006) study are novel because Jagg fluorescence was measured in the viable portion of the sperm population. Martinez-Pastor et al. (2004) observed some relationship between JC-1 staining and motility, although correlation with motility is regulated by many factors. A deeper study has been suggested by this research group. The mitochondrial stain 5,50,6,60-tetrachloro-1,10,3,30-tetraethylbenzimidazolyl-carbocyanine iodide (JC-1) does permit a distinction to be made between spermatozoa with poorly and highly functional mitochondria (Garner et al., 1997). In highly functional mitochondria, the concentration of JC-1 inside the mitochondria increases and the stain forms aggregates that fluoresce orange. When human spermatozoa were divided into high, moderate and low mitochondrial potential groups, based on JC-1 fluorescence, the in vitro fertilization rates were higher in the high potential group than in the low potential group (Kasai et al., 2002). JC-1 has also been used successfully to measure mitochondrial function using a fluorometer (Gravance et al., 2000). However, at greater concentrations, the probe aggregates and in the aggregate form fluoresces red-orange (Thomas et al., 1998).
Sperm DNA integrity is increasingly being recognized as an important measure of fertilizing efficiency that has better diagnostic and prognostic capabilities than standard sperm parameters like sperm morphology, concentration and motility. Routine semen parameters do not always reflect the quality of sperm DNA and early embryo development depends on the presence of normal DNA (Rodriguez-Martinez, 2007). Full sperm DNA integrity usually is defined as the absence of DNA nicks or single stranded (ss) breaks, double stranded (ds) breaks, and chemical modifications of the DNA. Of these, ds DNA breaks are the most mutagenic, because in the
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