The authors wish to acknowledge support from Dr. Lavoisier Ramos-Espiritu at the Rockefeller High Throughput and Spectroscopy Resource Center.

Sperm are collected from 8-16-week-old CD-1 male mice. Animal experiments were approved by Weill Cornell Medicine&#39;s Institutional Animal Care and Use Committee (IACUC).
1. Day prior to assay
<ol>
	<li>Preparation of sensor cartridge and extracellular flux analyzer calibrant
	<ol>
		<li>To hydrate the sensor cartridge, remove the sensor cartridge from the XFe96 Extracellular Flux Assay Kit and place the sensor cartridge upside down next to the utility plate.</li>
		<li>Fill a solution reservoir with 25 mL of double-distilled H2O using a multichannel pipette, add 200 &#181;L of H2O to each well of the utility plate. Place the sensor cartridge back into the utility plate and incubate overnight in a 37 &#176;C non-CO2 incubator. 
		NOTE: Cartridge needs to be hydrated for at least 4 h.</li>
		<li>Aliquot 25 mL of the extracellular flux analyzer calibrant into a 50 mL conical tube and incubate it overnight in a 37 &#176;C non-CO2 incubator.</li>
	</ol>
	</li>
	<li>Preparation of ConA-coated microplates
	<ol>
		<li>Dissolve 2.5 mg of ConA in 5 mL of double-distilled H2O to prepare a 0.5 mg/mL (w/v) stock.</li>
		<li>Using a multichannel pipette, fill each well of an extracellular flux analyzer 96-well plate with 50 &#181;L of the 0.5 mg/mL Con A solution. Leave the lid open and let the plate dry overnight at room temperature. 
		NOTE: Multiple plates can be coated at once and stored at 4 &#176;C for up to four weeks until ready for use.</li>
	</ol>
	</li>
	<li>Preparation of sperm buffer
	<ol>
		<li>Prepare 250 mL of extracellular flux analyzer TYH buffer with low HEPES containing 138 mM NaCl, 4.7 mM KCl, 1.7 mM CaCl2, 1.2 mM KH2PO4, 1.2 mM MgSO4, 5.6 mM glucose, and 1 mM HEPES. Heat the buffer to 37 &#176;C and adjust the pH to 7.4.</li>
	</ol>
	</li>
</ol>
2. Day of the assay
<ol>
	<li>Preparation for cartridge calibration
	<ol>
		<li>Replace the H2O in the utility plate with 200 &#181;L of XF calibrant per well and incubate for at least 1 h in a non-CO2 37 &#176;C incubator. 
		NOTE: Instead of calibrant, sterile-filtered PBS, pH 7.4 can be used.</li>
		<li>Heat 50 mL of TYH buffer at 37 &#176;C.</li>
	</ol>
	</li>
	<li>Generating a wave template for the extracellular flux analyzer
	<ol>
		<li>Turn the extracellular flux analyzer on and allow the temperature to stabilize to 37 &#176;C.</li>
		<li>Open the wave software and design a new template by opening a blank template (see Supplemental File 1, wave template).</li>
		<li>Open the Group definitions tab. Define assay media as TYH; pH 7.4 and cell type as mouse sperm, leave injection strategies and pretreatments blank. Create different groups for each assay condition; in this example, there are 6 different groups (TYH, TYH + db-cAMP/IBMX, 2-DG, 2-DG + db-cAMP/IBMX, Ant/Rot, Ant/Rot + db-cAMP/IBMX); dibutyryl cAMP (db-cAMP) and 3-Isobutyl-1-methylxanthine (IBMX) to induce capacitation, 2-deoxyglucose (2-DG) as glycolysis inhibitor, antimycin A (antA) and rotenone (rot) as inhibitor of complex III and complex I of the electron transport chain. 
		NOTE: To account for variability between wells, at least 7-8 wells per condition should be measured in parallel.</li>
		<li>Open the Plate map tab and define the plate map by assigning groups to specific wells. 
		NOTE: The four corner wells (A1, A12, H1, H12) will be filled with TYH buffer (no sperm) and will later be used for background subtraction.</li>
		<li>Open the Protocol tab, add 4 different measurement cycles, select the respective port and edit the measurement details (Figure 2, Table 1). Highlight measure after injection and do not highlight equilibrate.</li>
		<li>Save the assay template, fill out the project summary and define the saving location for the results file.</li>
	</ol>
	</li>
	<li>Preparation of compounds to load into sensor cartridge
	<ol>
		<li>Prepare 2 mL of 50 mM db-cAMP by dissolving 9.8 mg of compound in TYH buffer and add IBMX to a final concentration of 5 mM. 
		NOTE: IBMX has the tendency to precipitate after being diluted in TYH buffer. Precipitates can be dissolved by vortexing and incubating the TYH/db-cAMP/IBMX solution in a 37 &#176;C heat block for 5 min.</li>
		<li>Prepare 1 mL of 500 mM 2-DG by dissolving 82.1 mg of compound in TYH buffer.</li>
		<li>Prepare 1 mL of 5 &#181;M of AntA/Rot by diluting both drugs in TYH buffer. 
		NOTE: Compounds are diluted 10-fold by injection into the extracellular flux analyzer cartridge; therefore, be sure that injection solutions are 10x more concentrated.</li>
	</ol>
	</li>
	<li>Isolation of mouse sperm
	<ol>
		<li>Anesthetize 3 male mice between 8 and 16 weeks by isoflurane and sacrifice the mice by cervical dislocation after no response to toe pinching was detected. Remove the cauda epididymides and vasa deferentia.</li>
		<li>Place each cauda in 500 &#181;L of prewarmed TYH buffer in 24-well plate well, immobilize the cauda to the bottom of the plate using a pair of forceps and open the tissue by making 5-7 small incisions with feather scissors.</li>
		<li>Transfer the plate immediately into a non-CO2 37 &#176;C incubator and let the sperm disperse for 15 min.</li>
	</ol>
	</li>
	<li>Loading of the sensor cartridge
	<ol>
		<li>Two port-loading guides for port A and D and port B and C are provided in the Extracellular Flux kit. For loading a specific port, remove the lid from the sensor cartridge and align the letter of the respective port loading guide with the upper left corner of the cartridge. While injecting, use the fingertips of the non-injecting hand to hold the port loading guide in place and insert the pipette tips vertically into the port loading guide holes.</li>
		<li>Port A: Using a multichannel pipette, inject 20 &#181;L of the TYH buffer into every column.</li>
		<li>Port B: Inject 22 &#181;L of TYH buffer into column 1-4, inject 22 &#181;L of 500 mM 2-DG into column 5-8, and 25 &#181;L of 5 &#181;M AntA/Rot into column 9-12.</li>
		<li>Port C: Inject 25 &#181;L of TYH buffer into every column with odd numbers, inject 25 &#181;L of 10 mM db-cAMP/ 500 &#181;M IBMX into every column with even numbers.</li>
		<li>Place the sensor cartridge with the calibration plate into the extracellular flux analyzer and start the assay. Calibration takes 10 - 15 min.</li>
	</ol>
	</li>
	<li>Preparation of sperm plates
	<ol>
		<li>Dissolve 45 mg of BSA in 15 mL of TYH buffer (3 mg/mL w/v) and prepare three aliquots of 3 mL BSA/TYH solution each in 5 mL centrifuge tubes.</li>
		<li>Combine two sperm wells each in one 1.5 mL centrifuge tube, count the sperm using a hematocytometer and dilute the sperm to a concentration of 2 x 107 sperm/mL. 
		NOTE: Use cut pipette tips for pipetting sperm to avoid damaging the cells.</li>
		<li>Centrifuge the sperm for 3 min at 700 x g, remove the supernatant and add 1 mL of TYH buffer. Repeat the centrifugation step, remove the supernatant, and transfer each sperm suspension aliquot into one 5 mL centrifuge tube with 3 mL of BSA/TYH.</li>
		<li>Place 180 &#181;L of TYH buffer into each corner well (A1, A12, H1, H12) of a ConA-coated plate. Place 180 &#181;L of sperm suspension into each empty well of the ConA-coated plate (1.2 x 106 sperm/well).</li>
		<li>Centrifuge the sperm plate at 250 x g for 1 min, rotate the plate by 180&#176;, and centrifuge again at 250 x g for 1 min (lowest braking rate 1).</li>
		<li>Remove the calibration plate from the extracellular flux analyzer and add the sperm plate. Continue the assay.</li>
	</ol>
	</li>
	<li>Data extraction and analysis
	<ol>
		<li>After finishing the assay, remove the cartridge, open the results file, click the Export tab and export the completed run as a GraphPad Prism file (Supplemental File 2: Primary example data used for Figure 3).</li>
		<li>Duplicate the ECAR and OCR file by clicking on the New tab and then the Duplicate Family&#8230; tab (Figure 3A).</li>
		<li>Delete the first 7 rows of data (Figure 3B).</li>
		<li>Normalize to the data point before the cAMP/IBMX injection by baseline subtracting row 1. Click on the Analyze tab, then on the Remove baseline and column math tab. Highlight Selected row(s): First Row, Calculation: Ratio:Value/Baseline, and Subcolumns: Ignore Subcolumn. Click OK (Figure 3C).</li>
	</ol>
	</li>
</ol>

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Purification and characterization of two creatine kinase isozymes from sea urchin sperm.</a> Journal of Biological Chemistry. 262 (33), 16011-16019 (1987).</li><li>Gomendio, M., Tourmente, M., Roldan, E. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Why+mammalian+lineages+respond+differently+to+sexual+selection:+metabolic+rate+constrains+the+evolution+of+sperm+size.">Why mammalian lineages respond differently to sexual selection: metabolic rate constrains the evolution of sperm size.</a> Proceedings of the Royal Society of Biological Sciences. 278 (1721), 3135-3141 (2011).</li><li>Tourmente, M., Villar-Moya, P., Rial, E., Roldan, E. 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The authors have nothing to disclose.

The loss of sperm capacitation in the absence of certain metabolic substrates or critical metabolic enzymes revealed energy metabolism as a key factor supporting successful fertilization. A metabolic switch during cell activation is a well-established concept in other cell types, however, we are just beginning to understand how sperm adapt their metabolism to the increasing energy demand during capacitation. Using an extracellular flux analyzer, we developed an easily applicable tool to monitor changes in glycolysis and ...

The application of mass spectrometry has revolutionized the study of metabolism. Targeted metabolic profiling and metabolomic tracing allow precise monitoring of changes in energy metabolism. However, performing metabolomics successfully requires extensive training, experienced staff, and expensive, highly sensitive mass spectrometers not readily available to every laboratory. In recent years, using an extracellular flux analyzer, such as the Seahorse XFe96 has grown popular as a surrogate method for measuring changes in energy metabolism in various cell types1,2,3,4,5.
Sperm are highly specialized motile cells; whose task is to deliver the paternal genome to the oocyte. Sperm leaving the male reproductive tract after ejaculation are still functionally immature and cannot fertilize the oocyte because they are unable to penetrate the oocytes&#39; vestments. Sperm acquire fertilization competence as they transit through the female reproductive tract in a maturation process known as capacitation6,7. Freshly ejaculated sperm or sperm dissected from the cauda epididymis can be capacitated in vitro by incubation in defined capacitation media containing Ca2+, bicarbonate (HCO3-) or a cell-permeable cAMP analog (e.g., dibutyryl-cAMP), a cholesterol acceptor (e.g., bovine serum albumin, BSA), and an energy source (e.g., glucose). During capacitation, sperm modify their motility pattern into an asymmetric flagellar beat, representing a swimming mode called hyperactivation8,9, and they become competent to undergo the acrosome reaction7, where proteolytic enzymes are released that digest the oocytes&#39; vestments. These processes require energy, and similar to somatic cells, sperm generate ATP and other high energy compounds via glycolysis as well as mitochondrial TCA cycle and oxidative phosphorylation (oxphos)10. While multiple studies demonstrate that glycolysis is necessary and sufficient to support sperm capacitation11,12,13,14, the contribution of oxphos is less clear. Contrary to other cell types where glycolysis is physically coupled to the TCA cycle, sperm are highly compartmentalized and are thought to maintain these processes in separate flagellar compartments: the midpiece concentrates the mitochondrial machinery, whereas the key enzymes of glycolysis appear to be restricted to the principal piece15,16. This compartmentalization results in an ongoing debate about whether pyruvate produced in the principal piece by glycolysis can support mitochondrial oxphos in the midpiece, and whether ATP produced by oxphos in the midpiece would be able to diffuse sufficiently rapidly along the length of the flagellum to support the energy requirements in distal parts of the principal piece17,18,19. There is also support of a role for oxphos in sperm capacitation. Not only is oxphos more energetically favorable than glycolysis, generating 16 times more ATP than glycolysis, but midpiece volume and mitochondrial content are directly correlated with reproductive fitness in mammalian species which exhibit greater degrees of competition between males for mates20. Addressing these questions requires methods for examining the relative contributions of glycolysis and oxphos during sperm capacitation.
Tourmente et al. applied a 24-well extracellular flux analyzer to compare the energy metabolism of closely related mouse species with significantly different sperm performance parameters21. Instead of reporting the basal ECAR and OCR values of non-capacitated sperm, here, we adapt their method using a 96-well extracellular flux analyzer to monitor changes in energy metabolism during mouse sperm capacitation in real-time. We developed a method that allows simultaneously monitoring glycolysis and oxphos in real-time in sperm with beating flagella in up to twelve different experimental conditions by measuring the flux of oxygen (O2) and protons (H+) (Figure 1A). Due to the breakdown of pyruvate to lactate during glycolysis and the production of CO2 via the TCA-cycle, non-capacitated and capacitated sperm extrude H+ into the assay media which are detected by the extracellular flux analyzer via H+-sensitive fluorophores immobilized to the probe tip of a sensor cartridge. In parallel, O2 consumption by oxidative phosphorylation is detected via O2-sensitive fluorophores immobilized to the same probe tip (Figure 1B). Effective detection of the released H+ and consumed O2 requires a modified sperm buffer with low buffering capacity without bicarbonate or phenol red. Thus, to induce capacitation in the absence of bicarbonate, we adopted the use of a cell-permeable cAMP analog injected together with the broad-range PDE inhibitor IBMX22. Three additional independent injection ports allow the injection of pharmacological activators and/or inhibitors, which facilitates real-time detection of changes in cellular respiration and glycolysis rate due to experimental manipulation.

<table><tbody><tr><td>&lt;strong&gt;Reagents&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>2-Deoxy-D-glucose</td><td>Sigma-Aldrich</td><td>D8375</td><td>2-DG</td></tr><tr><td>3-Isobutyl-1-methylxanthine</td><td>Sigma-Aldrich</td><td>I7018</td><td>IBMX; prepare a 500 mM stock solution in DMSO (111.1 mg/ml) and store in small aliquots</td></tr><tr><td>Antimycin A</td><td>Sigma-Aldrich</td><td>A8674</td><td>AntA; prepare a 5 mM stock solution in DMSO (2.7 mg/ml) and store in small aliquots</td></tr><tr><td>Bovine serum albumin</td><td>Sigma-Aldrich</td><td>A1470</td><td>BSA</td></tr><tr><td>Calcium chloride</td><td>Sigma-Aldrich</td><td>C1016</td><td>CaCl2</td></tr><tr><td>Concanacalin A, Lectin from Arachis hypogaea (peanut)</td><td>Sigma-Aldrich</td><td>L7381</td><td>ConA</td></tr><tr><td>Glucose</td><td>Sigma-Aldrich</td><td>G7528</td><td></td></tr><tr><td>Hepes</td><td>Sigma-Aldrich</td><td>H0887</td><td></td></tr><tr><td>Isothesia</td><td>Henry Schein Animal Health</td><td>1169567761</td><td>Isoflurane</td></tr><tr><td>Magnesium sulfate</td><td>Sigma-Aldrich</td><td>M2643</td><td>MgSO4</td></tr><tr><td>N6,2&#039;-O-Dibutyryladenosine 3&#039;,5&#039;-cyclic monophosphate sodium salt</td><td>Sigma-Aldrich</td><td>D0627</td><td>db-cAMP</td></tr><tr><td>Potassium chloride</td><td>Sigma-Aldrich</td><td>P9333</td><td>KCl</td></tr><tr><td>Potassium dihydrogen phosphate</td><td>Sigma-Aldrich</td><td>P5655</td><td>KH2PO4</td></tr><tr><td>Rotenone</td><td>Cayman Chemical Company</td><td>13995</td><td>Rot; prepare a 5 mM stock solution in DMSO (2mg/ml) and store in small aliquots</td></tr><tr><td>Sodium bicarbonate</td><td>Sigma-Aldrich</td><td>S5761</td><td>NaHCO3-</td></tr><tr><td>Sodium chloride</td><td>Sigma-Aldrich</td><td>S9888</td><td>NaCl</td></tr><tr><td>&lt;strong&gt;Equipment and materials&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>12 channel pipette 10-100 &amp;mu;L</td><td>eppendorf</td><td>ES-12-100</td><td></td></tr><tr><td>12 channel pipette 50-300 &amp;mu;L</td><td>vwr</td><td>613-5257</td><td></td></tr><tr><td>37 &amp;deg;C, non-CO&lt;sub&gt;2&lt;/sub&gt; incubator</td><td>vwr</td><td>1545</td><td></td></tr><tr><td>5 mL cetrifuge tubes</td><td>eppendorf</td><td>30119380</td><td></td></tr><tr><td>50 mL conical centrifuge tubes</td><td>vwr</td><td>76211-286</td><td></td></tr><tr><td>Centrifuge with plate adapter</td><td>Thermo Scientific</td><td>IEC FL40R</td><td></td></tr><tr><td>Dissection kit</td><td>World Precision Instruments</td><td>MOUSEKIT</td><td></td></tr><tr><td>Inverted phase contrast microscope with 40X objective</td><td>Nikon</td><td></td><td></td></tr><tr><td>OctaPool Solution Reservoirs, 25 ml, divided</td><td>Thomas Scientific</td><td>1159X93</td><td></td></tr><tr><td>OctaPool Solution Reservoirs, 25 mL, divided</td><td>Thomas Scientific</td><td>1159X95</td><td></td></tr><tr><td>Seahorse XFe96 Analyzer</td><td>Agilent</td><td></td><td></td></tr><tr><td>Seahorse XFe96 FluxPak</td><td>Agilent</td><td>102416-100</td><td>Also sold as XFe96 FluxPak mini (102601-100) with 6 instead of 18 cartidges.</td></tr></tbody></table>

using an extracellular flux analyzer to measure changes in glycolysis and oxidative phosphorylation during mouse sperm capacitation

Mammalian sperm acquire fertilization capacity in the female reproductive tract in a process known as capacitation. Capacitation-associated processes require energy. There remains an ongoing debate about the sources generating the ATP which fuels sperm progressive motility, capacitation, hyperactivation, and acrosome reaction. Here, we describe the application of an extracellular flux analyzer as a tool to analyze changes in energy metabolism during mouse sperm capacitation. Using H+- and O2- sensitive fluorophores, this method allows monitoring glycolysis and oxidative phosphorylation in real-time in non-capacitated versus capacitating sperm. Using this assay in the presence of different energy substrates and/or pharmacological activators and/or inhibitors can provide important insights into the contribution of different metabolic pathways and the intersection between signaling cascades and metabolism during sperm capacitation.

This method uses an extracellular flux analyzer to monitor real-time changes in the rate of glycolysis and oxphos during mouse sperm capacitation. Figure 4 shows an exemplary experiment where sperm were capacitated in the presence of glucose as the only energy substrate and 2-DG and antimycin and rotenone as pharmacological modulators. The energy substrate in the extracellular flux analyzer TYH buffer and the pharmacological modulators can be freely selected ...

Watch this Scientific Journal Video about Using an Extracellular Flux Analyzer to Measure Changes in Glycolysis and Oxidative Phosphorylation during Mouse Sperm Capacitation at JoVE.com

Using an Extracellular Flux Analyzer to Measure Changes in Glycolysis and Oxidative Phosphorylation during Mouse Sperm Capacitation

We describe the application of an extracellular flux analyzer to monitor real-time changes in glycolysis and oxidative phosphorylation during mouse sperm capacitation.

Mammalian sperm acquire fertilization capacity in the female reproductive tract in a process known as capacitation. Capacitation-associated processes ...

using-an-extracellular-flux-analyzer-to-measure-changes-glycolysis

Universidad de Cartagena UDEC

Research

JoVE Journal

Biochemistry

7.8K Views.  Weill Cornell Medical College. We describe the application of an extracellular flux analyzer to monitor real-time changes in glycolysis and oxidative phosphorylation during mouse sperm capacitation.

Video: Using an Extracellular Flux Analyzer to Measure Changes in Glycolysis and Oxidative Phosphorylation during Mouse Sperm Capacitation

Exploring Mitochondrial Energy Metabolism of Single 3D Microtissue Spheroids Using Extracellular Flux Analysis

Three-dimensional (3D) cellular aggregates, termed spheroids, have become the forefront of in vitro cell culture in recent years. In contrast to culturing cells as two-dimensional, single-cell monolayers (2D culture), spheroid cell culture promotes, regulates, and supports physiological cellular architecture and characteristics that exist in vivo, including the expression of extracellular matrix proteins, cell signaling, gene expression, protein production, differentiation, and proliferation. The importance of 3D culture has been recognized in many research fields, including oncology, diabetes, stem cell biology, and tissue engineering. Over the last decade, improved methods have been developed to produce spheroids and assess their metabolic function and fate.
Extracellular flux (XF) analyzers have been used to explore mitochondrial function in 3D microtissues such as spheroids using either an XF24 islet capture plate or an XFe96 spheroid microplate. However, distinct protocols and the optimization of probing mitochondrial energy metabolism in spheroids using XF technology have not been described in detail. This paper provides detailed protocols for probing mitochondrial energy metabolism in single 3D spheroids using spheroid microplates with the XFe96 XF analyzer. Using different cancer cell lines, XF technology is demonstrated to be capable of distinguishing between cellular respiration in 3D spheroids of not only different sizes but also different volumes, cell numbers,&#160;DNA content and type.
The optimal mitochondrial effector compound concentrations of oligomycin, BAM15, rotenone, and antimycin A are used to probe specific parameters of mitochondrial energy metabolism in 3D spheroids. This paper also discusses methods to normalize data obtained from spheroids and addresses many considerations that should be considered when exploring spheroid metabolism using XF technology. This protocol will help drive research in advanced in vitro spheroid models.

These protocols will help users probe mitochondrial energy metabolism in 3D cancer cell-line-derived spheroids using Seahorse extracellular flux analysis.

Three-dimensional (3D) cellular aggregates, termed spheroids, have become the forefront of in vitro cell culture in recent years. In contrast to culturing ...

Cancer Research

Fluorimetric Techniques for the Assessment of Sperm Membranes

Standard spermiograms describing sperm quality are mostly based on the physiological and visual parameters, such as ejaculate volume and concentration, motility and progressive motility, and sperm morphology and viability. However, none of these assessments is good enough to predict the semen quality. Given that maintenance of sperm viability and fertilization potential depends on membrane integrity and intracellular functionality, evaluation of these parameters might enable a better prediction of sperm fertilization competence. Here, we describe three feasible methods to evaluate sperm quality using specific fluorescent probes combined with fluorescence microscopy or flow cytometry analyses. Analyses assessed plasma membrane integrity using 4&#39;,6-diamidino-2-phenylindole (DAPI) and propidium iodide (PI), acrosomal membrane integrity using fluorescein isothiocyanate-conjugated Pisum sativum agglutinin (FITC-PSA) and mitochondrial membrane integrity using 5,5&#39;,6,6&#39;-tetra-chloro-1,1&#39;,3,3&#39;-tetraethylbenzimidazolyl carbocyanine iodide (JC-1). Combinations of these methods are also presented. For instance, use of annexin V combined with PI fluorochromes enables assessing apoptosis and calculating the proportion of apoptotic sperm (apoptotic index). We believe that these methodologies, which are based on examining spermatozoon membranes, are very useful for the evaluation of sperm quality.

Here, we present methodologies to evaluate spermatozoan membrane integrity, a cellular feature associated with sperm fertilization competence. We describe three techniques for the fluorimetric assessment of sperm membranes: simultaneous staining with specific fluorescent probes, fluorescence microscopy, and advanced sperm-dedicated flow cytometry. Examples of combining the methodologies are also presented.

Standard spermiograms describing sperm quality are mostly based on the physiological and visual parameters, such as ejaculate volume and concentration, ...

Developmental Biology

Combining Imaging with a Stimulant Delivery System to Study Live Mouse Sperm

Real-Time Imaging of Acrosomal Calcium Dynamics and Exocytosis in Live Mouse Sperm

Acrosome exocytosis (AE), in which the sperm&#39;s single exocytotic vesicle fuses with the plasma membrane, is a complex, calcium-dependent process essential for fertilization. However, our understanding of how calcium signaling regulates AE is still incomplete. In particular, the interplay between intra-acrosomal calcium dynamics and the intermediate steps leading to AE is not well-defined. Here, we describe a method that provides spatial and temporal insights into acrosomal calcium dynamics and their relationship to membrane fusion and subsequent exocytosis of the acrosome vesicle. The method utilizes a novel transgenic mouse expressing an Acrosome-targeted Sensor for Exocytosis (AcroSensE). The sensor combines a genetically encoded calcium indicator (GCaMP) fused with mCherry. This fusion protein was specifically designed to enable the concurrent observation of acrosomal calcium dynamics and membrane fusion events. Real-time monitoring of acrosomal calcium dynamics and AE in live AcroSensE sperm is achieved using a combination of high frame-rate imaging and a stimulant delivery system that can target single sperm. This protocol also provides several examples of basic methods to quantify and analyze the raw data. Because the AcroSensE model is genetically encoded, its scientific significance can be augmented by using readily available genetic tools, such as crossbreeding with other mouse genetic models or gene-editing (CRISPR) based methods. With this strategy, the roles of additional signaling pathways in sperm capacitation and fertilization can be resolved. In summary, the method described here provides a convenient and effective tool to study calcium dynamics in a specific subcellular compartment-the sperm acrosome-and how those dynamics regulate the intermediate steps leading to membrane fusion and acrosome exocytosis.

Author Spotlight: A Live Cell Imaging Technique to Study Calcium Signaling and Acrosome Exocytosis in Mouse Sperm 

The AcroSensE mouse model and live cell imaging methods described here provide a new approach to studying calcium dynamics in the subcellular compartment of the sperm acrosome and how they regulate intermediate steps leading to membrane fusion and acrosome exocytosis.

Acrosome exocytosis (AE), in which the sperm&#39;s single exocytotic vesicle fuses with the plasma membrane, is a complex, calcium-dependent process ...

Measuring Intracellular Ca2+ Changes in Human Sperm using Four Techniques: Conventional Fluorometry, Stopped Flow Fluorometry, Flow Cytometry and Single Cell Imaging

Spermatozoa are male reproductive cells especially designed to reach, recognize and fuse with the egg. To perform these tasks, sperm cells must be prepared to face a constantly changing environment and to overcome several physical barriers. Being in essence transcriptionally and translationally silent, these motile cells rely profoundly on diverse signaling mechanisms to orient themselves and swim in a directed fashion, and to contend with challenging environmental conditions during their journey to find the egg. In particular, Ca2+-mediated signaling is pivotal for several sperm functions: activation of motility, capacitation (a complex process that prepares sperm for the acrosome reaction) and the acrosome reaction (an exocytotic event that allows sperm-egg fusion). The use of fluorescent dyes to track intracellular fluctuations of this ion is of remarkable importance due to their ease of application, sensitivity, and versatility of detection. Using one single dye-loading protocol we utilize four different fluorometric techniques to monitor sperm Ca2+ dynamics. Each technique provides distinct information that enables spatial and/or temporal resolution, generating data both at single cell and cell population levels.

Measuring Intracellular Ca2+ Changes in Human Sperm using Four Techniques: Conventional Fluorometry, Stopped Flow Fluorometry, Flow Cytometry and Single Cell Imaging

Intracellular Ca2+ dynamics are very important in sperm physiology and Ca2+-sensitive fluorescent dyes constitute a versatile tool to study them. Population experiments (fluorometry and stopped flow fluorometry) and single cell experiments (flow cytometry and single cell imaging) are used to track spatio-temporal [Ca2+] changes in human sperm cells.

Spermatozoa  are male reproductive cells especially designed to reach, recognize and fuse  with the egg. To perform these tasks, sperm cells must be ...

Biology

Analyzing Oxygen Consumption Rate in Primary Cultured Mouse Neonatal Cardiomyocytes Using an Extracellular Flux Analyzer

Mitochondria and oxidative metabolism are critical for maintaining cardiac muscle function. Research has shown that mitochondrial dysfunction is an important contributing factor to impaired cardiac function found in heart failure. By contrast, restoring defective mitochondrial function may have beneficial effects to improve cardiac function in the failing heart. Therefore, studying the regulatory mechanisms and identifying novel regulators for mitochondrial function could provide insight which could be used to develop new therapeutic targets for treating heart disease. Here, cardiac myocyte mitochondrial respiration is analyzed using a unique cell culture system. First, a protocol has been optimized to rapidly isolate and culture high viability neonatal mouse cardiomyocytes. Then, a 96-well format extracellular flux analyzer is used to assess the oxygen consumption rate of these cardiomyocytes. For this protocol, we optimized seeding conditions and demonstrated that neonatal mouse cardiomyocytes oxygen consumption rate can be easily assessed in an extracellular flux analyzer. Finally, we note that our protocol can be applied to a larger culture size and other studies, such as intracellular signaling and contractile function analysis.

The goal of this protocol is to illustrate how to use mouse neonatal cardiomyocytes as a model system to examine how various factors can alter oxygen consumption in the heart.

Mitochondria and oxidative metabolism are critical for maintaining cardiac muscle function. Research has shown that mitochondrial dysfunction is an ...

Medicine

Real-Time Measurement of the Mitochondrial Bioenergetic Profile of Neutrophils

Neutrophils are the first line of defense and the most abundant leukocytes in humans. These effector cells perform functions such as phagocytosis and oxidative burst, and create neutrophil extracellular traps (NETs) for microbial clearance. New insights into the metabolic activities of neutrophils challenge the early concept that they primarily rely on glycolysis. Precise measurement of metabolic activities can unfold different metabolic requirements of neutrophils, including the tricarboxylic acid (TCA) cycle (also known as the Krebs cycle), oxidative phosphorylation (OXPHOS), pentose phosphate pathway (PPP), and fatty acid oxidation (FAO) under physiological conditions and in disease states. This paper describes a step-by-step protocol and prerequirements to measure oxygen consumption rate (OCR) as an indicator of mitochondrial respiration on mouse bone marrow-derived neutrophils, human blood-derived neutrophils, and the neutrophil-like HL60 cell line, using metabolic flux analysis on a metabolic extracellular flux analyzer. This method can be used for quantifying the mitochondrial functions of neutrophils under normal and disease conditions.

We describe stepwise protocols measuring the mitochondrial respiration of mouse and human neutrophils and HL60 cells using the metabolic extracellular flux analyzer.

Neutrophils are the first line of defense and the most abundant leukocytes in humans. These effector cells perform functions such as phagocytosis and ...

Immunology and Infection

Measurement and Analysis of Extracellular Acid Production to Determine Glycolytic Rate

Extracellular measurement of oxygen consumption and acid production is a simple and powerful way to monitor rates of respiration and glycolysis1. Both mitochondrial (respiration) and non-mitochondrial (other redox) reactions consume oxygen, but these reactions can be easily distinguished by chemical inhibition of mitochondrial respiration. However, while mitochondrial oxygen consumption is an unambiguous and direct measurement of respiration rate2, the same is not true for extracellular acid production and its relationship to glycolytic rate 3-6. Extracellular acid produced by cells is derived from both lactate, produced by anaerobic glycolysis, and CO2, produced in the citric acid cycle during respiration. For glycolysis, the conversion of glucose to lactate- + H+ and the export of products into the assay medium is the source of glycolytic acidification. For respiration, the export of CO2, hydration to H2CO3 and dissociation to HCO3- + H+ is the source of respiratory acidification. The proportions of glycolytic and respiratory acidification depend on the experimental conditions, including cell type and substrate(s) provided, and can range from nearly 100% glycolytic acidification to nearly 100% respiratory acidification 6. Here, we demonstrate the data collection and calculation methods needed to determine respiratory and glycolytic contributions to total extracellular acidification by whole cells in culture using C2C12 myoblast cells as a model.

Glycolysis is a defining metabolic marker in multiple biological systems. Monitoring glycolysis by measuring the extracellular flux of H+ is common, but requires correction to be quantitative and unambiguous. Here, we demonstrate how to gather and correct extracellular flux data to distinguish between respiratory and glycolytic sources of extracellular acidification.

Extracellular measurement of oxygen consumption and acid production is a simple and powerful way to monitor rates of respiration and glycolysis1. Both ...

Determining Respiration in Human Sperm Cells by High-Resolution Respirometry

High-Resolution Respirometry to Assess Mitochondrial Function in Human Spermatozoa

Semen quality is often studied by routine semen analysis, which is descriptive and often inconclusive. Male infertility is associated with altered sperm mitochondrial activity, so the measurement of sperm mitochondrial function is an indicator of sperm quality. High-resolution respirometry is a method of measuring the oxygen consumption of cells or tissues in a closed-chamber system. This technique can be implemented to measure respiration in human sperm and provides information about the quality and integrity of the sperm mitochondria. High-resolution respirometry allows the cells to move freely, which is an a priori advantage in the case of sperm. This technique can be applied with intact or permeabilized spermatozoa and allows for the study of intact sperm mitochondrial function and the activity of individual respiratory chain complexes. The high-resolution oxygraph instrument uses sensors to measure the oxygen concentration coupled with sensitive software to calculate the oxygen consumption. The data are used to calculate respiratory indices based on the oxygen consumption ratios. Consequently, the indices are the proportions of two oxygen consumption rates and are internally normalized to the cell number or protein mass. The respiratory indices are an indicator of sperm mitochondrial function and dysfunction.

Author Spotlight: Advancing Male Infertility Research by Unraveling Sperm Metabolism and Mitochondrial Function 

The analysis of sperm mitochondrial function by high-resolution respirometry permits the measurement of the oxygen consumption of freely moving spermatozoa in a closed-chamber system. The technique can be applied to measure respiration in human spermatozoa, which provides information on sperm mitochondrial characteristics and integrity.

Semen quality is often studied by routine semen analysis, which is descriptive and often inconclusive. Male infertility is associated with altered sperm ...

Measurement of Energy Metabolism in Explanted Retinal Tissue Using Extracellular Flux Analysis

High acuity vision is a heavily energy-consuming process, and the retina has developed several unique adaptations to precisely meet such demands while maintaining transparency of the visual axis. Perturbations to this delicate balance cause blinding illnesses, such as diabetic retinopathy. Therefore, the understanding of energy metabolism changes in the retina during disease is imperative to the development of rational therapies for various causes of vison loss. The recent advent of commercially-available extracellular flux analyzers has made the study of retinal energy metabolism more accessible. This protocol describes the use of such an analyzer to measure contributions to retinal energy supply through its two principle arms - oxidative phosphorylation and glycolysis - by quantifying changes in oxygen consumption rates (OCR) and extracellular acidification rates (ECAR) as proxies for these pathways. This technique is readily performed in explanted retinal tissue, facilitating assessment of responses to multiple pharmacologic agents in a single experiment. Metabolic signatures in retinas from animals lacking rod photoreceptor signaling are compared to wild-type controls using this method. A major limitation in this technique is the lack of ability to discriminate between light-adapted and dark-adapted energy utilization, an important physiologic consideration in retinal tissue.

This technique describes real time recording of oxygen consumption and extracellular acidification rates in explanted mouse retinal tissues using an extracellular flux analyzer.

High acuity vision is a heavily energy-consuming process, and the retina has developed several unique adaptations to precisely meet such demands while ...

Assessment of Cellular Bioenergetics in Mouse Hematopoietic Stem and Primitive Progenitor Cells using the Extracellular Flux Analyzer

Under steady state, hematopoietic stem cells (HSCs) remain largely quiescent and are believed to be predominantly reliant on glycolysis to meet their energetic needs. However, under stress conditions such as infection or blood loss, HSCs become proliferative and rapidly produce downstream progenitor cells, which in turn further differentiate, ultimately producing mature blood cells. During this transition and differentiation process, HSCs exit from quiescence and rapidly undergo a metabolic switch from glycolysis to oxidative phosphorylation (OxPHOS). Various stress conditions, such as aging, cancer, diabetes, and obesity, can negatively impact mitochondrial function and thus can alter the metabolic reprogramming and differentiation of HSCs and progenitors during hematopoiesis. Valuable insights into glycolytic and mitochondrial functions of HSCs and progenitors under normal and stress conditions can be gained through the assessment of their extracellular acidification rate (ECAR) and oxygen consumption rate (OCR), which are indicators of cellular glycolysis and mitochondrial respiration, respectively. 
Here, a detailed protocol is provided to measure ECAR and OCR in mouse bone marrow-derived lineage-negative cell populations, which include both hematopoietic stem and primitive progenitor cells (HSPCs), using the extracellular flux analyzer. This protocol describes approaches to isolate lineage-negative cells from mouse bone marrow, explains optimization of cell seeding density and concentrations of 2-deoxy-D-glucose (2-DG, a glucose analog that inhibits glycolysis) and various OxPHOS-targeted drugs (oligomycin, FCCP, rotenone, and antimycin A) used in these assays, and describes drug treatment strategies. Key parameters of glycolytic flux, such as glycolysis, glycolytic capacity, and glycolytic reserve, and OxPHOS parameters, such as basal respiration, maximal respiration, proton leak, ATP production, spare respiratory capacity, and coupling efficiency, can be measured in these assays. This protocol allows ECAR and OCR measurements on non-adherent HSPCs and can be generalized to optimize analysis conditions for any type of suspension cells.

The method presented here summarizes optimized protocols for assessing cellular bioenergetics in non-adherent mouse hematopoietic stem and primitive progenitor cells (HSPCs) using the extracellular flux analyzer to measure the extracellular acidification rate (ECAR) and oxygen consumption rate (OCR) of HSPCs in real time.

Under steady state, hematopoietic stem cells (HSCs) remain largely quiescent and are believed to be predominantly reliant on glycolysis to meet their ...