This work was funded in part by the National Science Foundation grant CHE-160944 to Mary E. Konkle and Michael A. Menze.

1. Cell Culture
<ol>
	<li>Maintain human hepatocellular carcinoma (HepG2) cells in Dulbecco&#39;s Modified Eagle&#39;s medium (DMEM) containing 10% fetal bovine serum (FBS) and additional substrates (10 mM glucose, 2 mM glutamine, and 1 mM pyruvate) at 37 &#176;C in a cell culture incubator (5% CO2, 95% air, 100% humidity). 
	NOTE: Use the above DMEM formulation when DMEM is referenced in later steps for respirometry and calorimetry.</li>
	<li>Plate the cells at 1 x 106 cells per 100 mm culture plate and subculture them every 7 days or before reaching 90% confluency and renew the medium every 3 - 4 days.</li>
	<li>Perform the experiments when cells are 60 - 80% confluent.</li>
</ol>
2. Preparation of Respirometer and Calorimeter
<ol>
	<li>Pipet 2.2 mL of DMEM into the respirometer chamber, insert the stopper fully, and adjust it with the stopper spacer tool to allow for optimal oxygen diffusion from air to medium.</li>
	<li>Stir continuously at 37 &#176;C until oxygen concentration (blue trace) and oxygen flux (red trace) are stable. Ensure the respirometer&#39;s software is programmed to account for the oxygen solubility of the medium used in the experiment. 
	NOTE: Different media have different oxygen solubility coefficients (e.g., DMEM = 0.92).</li>
	<li>After stabilization of the oxygen concentration in DMEM with the chamber open, select a stable portion of the oxygen concentration trace and calibrate it for 100% air saturation.</li>
	<li>Calibrate for 0% oxygen selecting a stable portion of the oxygen concentration trace when no oxygen is present in the chamber. Perform this step after 20 &#181;L titration of 230 mM sodium dithionite or after all oxygen is consumed by the biological sample at the end of an experiment.</li>
	<li>After the 100% air calibration of the respirometer, close the stopper and ensure that no bubbles are present in the chamber. 
	NOTE: a slight increase in oxygen flux will be detected in the closed chamber as the polarographic oxygen sensor (POS) consumes oxygen in order to measure the oxygen partial pressure.</li>
	<li>After ~10 min of the stopper being closed, confirm that the respirometer is ready for HepG2 cells by observing a stable oxygen flux. 
	NOTE: The calorimeter used here utilizes stainless steel ampules and requires one ampule to be used as a reference.</li>
	<li>Add 2.5 mL of H2O (dH2O) [equal volume as the samples (step 4)] to the reference ampule of the calorimeter and tighten the cap, using pliers if necessary. Clean the sealed ampule with airflow and slowly lower the ampule to position 1 in the reference channel of the calorimeter. Remain at position 1 until biological sample is prepared.</li>
</ol>
3. Preparation of HepG2 Cells for Respirometry and Calorimetry
<ol>
	<li>Confirm that the respirometer and calorimeter are prepared and calibrated. Warm DMEM and trypsin to 37 &#176;C in a water bath.</li>
	<li>Add DMEM to a fresh 100-mm plate and place in the incubator to allow for the equilibration of the medium for both CO2 and temperature. 
	NOTE: This medium will be used for the calorimetry experiment, so the appropriate volume depends on the number of ampules that will be used.</li>
	<li>Confirm with an inverted light microscope that the cells are at 60 - 80% confluent, then wash the 100-mm plate 2x with 10 mL of room temperature phosphate-buffered saline (PBS).</li>
	<li>Add 3 mL of the 37-&#176;C trypsin to the 100-mm plate of cells and incubate them for 7 - 10 min at 37 &#176;C in the cell culture incubator. Neutralize the trypsin with 3 mL of 37-&#176;C DMEM and suspend by gently pipetting it ~20 times with a 5 mL pipet.</li>
	<li>Transfer the 6 mL of resuspended cells in DMEM and trypsin into a sterile 15-mL conical tube and centrifuge it for 5 min at 175 x g. Remove the liquid without disturbing the cell pellet.</li>
	<li>Resuspend the cell pellet in ~5 mL of DMEM and pipet it thoroughly to ensure the cells are sufficiently dissociated from one another.</li>
	<li>Remove ~100 &#181;L of the well-mixed sample and perform a thorough cell count utilizing all 8 fields on a hemocytometer to determine the total number of cells. Then calculate the volume for resuspension of the cells in order to generate ~2 x 106 cells per 50 &#181;L. 
	NOTE: This will ensure that 2 x 106 cells are added to each chamber of the respirometer with minimal medium displacement.</li>
	<li>Centrifuge the cells for 5 min at 175 x g. Resuspend the pellet in the calculated volume of DMEM from step 3.7.</li>
	<li>Perform a cell count to determine the volume required to inject 2 x 106 cells per chamber in the respirometer (this should be ~50 &#181;L). Store the cells for the respirometric measurement on ice until ready.</li>
	<li>While the concentrated cell sample is on ice, determine the dilution required to produce a concentration of 1 x 105 cells/mL for the calorimetry.</li>
	<li>Mix the sample well and prepare ~3 mL of the cells at 1 x 105 cells/mL in DMEM for each ampule. 
	NOTE: The DMEM used for the sample dilution should be the equilibrated medium prepared in step 3.3.</li>
</ol>
4. Calorimetry
<ol>
	<li>Ensure that calorimeter is connected to the computer and the calorimetric signal in &#181;W is observable and stable.</li>
	<li>Add 2.5 mL of the cells at 1 x 105 cells/mL (~2.5 x 105 cells) to each ampule, ensuring sufficient air is between the lid of the ampule and the medium to allow for gas diffusion.</li>
	<li>Immediately tighten the cap, using pliers if necessary. Clean the sealed ampule, using airflow to remove any external debris, and slowly lower the ampule to position 1 of the calorimeter. Record the time the ampule is lowered to position 1.</li>
	<li>Allow the ampule to sit at position 1 for 15 min. 
	NOTE: During this 15 min period, cells are to be injected into the respirometer (step 5). This ensures that the same interval of time is used for both heat production and oxygen consumption when the CR ratio is determined.</li>
	<li>After 15 min at position 1, slowly lower both the reference and sample ampules to position 2. Record the time the ampules were lowered and measure for at least 30 min.</li>
</ol>
5. Respirometry
<ol>
	<li>During the 15-min interval in which the ampule is at position 1 in the calorimeter, begin the respirometric studies.</li>
	<li>Thoroughly mix the cell sample by pipetting to ensure a homogenous solution and inject &#60;50 &#181;L of the cell sample into each chamber of the respirometer for a final concentration of ~2 x 106 cells/chamber. Record the time the cells are injected into the respirometer.</li>
	<li>After the injection, the HepG2 cells are continuously stirred at 37 &#176;C. Wait at least 20 - 30 min for both a stabilization of the oxygen flux and a steady decrease in the oxygen concentration.</li>
	<li>Select the O2 flux per V tab on the right of the screen and highlight a stable portion of oxygen flux. Select marks, then statistics and record data for O2 flux per V. This recording will be used in calculation of the CR ratio. 
	NOTE: Steps 5.5 and 5.6 are optional. Leak respiration and maximal uncoupled respiration will be revealed by titrations of oligomycin and FCCP.</li>
	<li>Inject 1 &#181;L of 4mg/mL oligomycin into each chamber and allow ~ 10 min for oxygen flux stabilization. Record stable leak respiration rate by following steps performed in 5.4 by highlighting a stable portion of the oxygen flux trace after oligomycin addition.</li>
	<li>Titrate 2-&#181;L injections of 0.2 mM FCCP into each chamber, allowing for oxygen flux stabilization after each injection. Continue the titrations until the maximal flux is reached, as indicated by a slight decline in the subsequent FCCP titration. Record the stable respiration rate during the maximal flux.</li>
</ol>
6. Calculation of Calorespirometric Ratio
<ol>
	<li>Perform a final cell count of the samples prepared at ~ 1 x 105 cells/mL utilizing all 8 fields of a hemocytometer to determine the total number of cells added to the ampule (2.5 mL).</li>
	<li>Determine a time to record measurements from both calorimeter and respirometer at which&#160;stable signals are&#160;present at identical times. Reference the time recorded when the ampule was lowered to position 2 and the time of injection of cells into the respirometer.</li>
	<li>Record the heat production by placing the cursor over the trace displaying heat output for the respected ampule at the time determined in step and express as &#181;W/million cells. Collect oxygen consumption data and ensure that it is expressed as pmol O2 x&#160;s-1&#160;x&#160;million cells.</li>
	<li>To calculate the CR ratio, first convert &#181;W to kJ/s, and then divide kJ/s over mol of O2/s to obtain the CR ratio expressed as kJ/mol O2. NOTE: The CR ratio is presented in kJ/mol O2. 
	(see&#160;<a href="/files/ftp_upload/57724/JoVE_Supplemental_File.docx" target="_blank">Supplementary FIle 1</a>)</li>
</ol>

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The authors have nothing to disclose.

The objective of calorespirometry is to quantitatively evaluate the contributions of aerobic and anaerobic pathways to metabolic activity and obtain a composite view of cellular energy flux. This is accomplished by a simultaneous measurement of heat dissipation and oxygen flux followed by a comparison of the calculated CR ratio with the theoretical oxycaloric equivalent. Several critical steps must be considered for reproducible and reliable data. The maintenance of a sterile, healthy cell culture is critical. Contamination by bacteria or fungi may result in misappropriated heat production or oxygen consumption, rendering the CR ratio meaningless. Further, improper cell counts, poor sample mixing, or clumped cells can also result in an inaccurate CR ratio. A critical step in regard to the respirometer is proper sample preparation so that the injection is below 50 &#181;L in volume. This volume ensures a minimal amount of medium is displaced from the chamber and the oxygen consumption is correctly attributed to 2 x 106 cells. Once routine respiration is observed, the proposed protocol is compatible with two additional titrations (oligomycin and FCCP), which illuminate leak respiration and ETS capacity of the mitochondria. For this application, excessive addition of FCCP can inhibit respiration and prevent&#160;maximal flux; therefore, at least 3 - 5 titrations should be performed prior to reaching the maximal uncoupled respiration.
Maintenance and proper calibration of both the respirometer and calorimeter is critical for calorespirometry. The titration protocol for background oxygen consumption is highly recommended and is performed with step-wise titrations of dithionite to reach decreased oxygen levels within the chamber. This is an essential measure to confirm tightly sealed chambers and correct for oxygen back diffusion into the chambers, especially if measurements are performed at a low oxygen tension. Also, performing a stirrer test before addition of cells can provide information about the responsiveness of the POS. If responsiveness is poor or the oxygen flux is not stable (Figure 2), service to the POS is required. Improper cleaning of the respirometer chambers can leave behind residual amounts of inhibitors or uncouplers, which may affect subsequent experiments. Air bubbles in the sealed chamber are also capable of interfering with the measurements and can be introduced during the closing of the chamber or during sample injection if air bubbles are present within the syringe. The calorimeter&#8217;s protocol is more direct, but it is also susceptible to experimental errors. For example, the improper sealing of the ampule will allow evaporation to occur which massively changes heat flow.
One of the limitations of this method is that not all cells can be cultured in suspension and many cell lines are adherent to the cell culture plate or substrate. In order to perform experiments, attached cells have to be enzymatically dislodged from the plate through the use of dissociation agents such as trypsin. Cells in this state may not be in the logarithmic growth phase and their physiology may be altered. Thus,&#160;subsequent analysis via the calorimeter and respirometer of recently trypsinized cells may assess disrupted cellular activities. Further, a limitation in the presented calorimetric protocol is the sedimentation of cells in the enclosed ampules, which may lead to local hypoxia around the cells due to slow oxygen diffusion in aqueous solutions. We have determined that ~3 x 105 cells is the upper limit compatible with the ampules used in this protocol. Increasing the number of cells can generate a hypoxic environment from the sedimentation of the cells, compromising the results. Thus, optimization is critical if a different cell line is investigated and alternative approaches should be considered such as increasing the medium density to reduce the sedimentation of cells24. Ampules are available that permit stirring; however, it is critical to ensure a minimal signal-to- noise ratio is present during the stirring. Additionally, calorimeters besides the one employed in this protocol are compatible with calorespirometry and their capabilities may be beyond what we have presented. Another limitation of calorespirometric analysis via a two-chamber respirometer is the inability for a high-throughput analysis, which would be most relevant for pharmaceutical development. This limitation, however, is offset by the capacity for a high-resolution characterization of the compound&#39;s influence on the respiratory chain and mitochondrial physiology.
The capacity for both high-resolution and simultaneous measurement of heat output and oxygen flux from the same sample is a significant strength of this method. A potential gold standard when using adherent cells would be to both culture and analyze cells grown on microcarrier beads in suspension. This method would avoid the negative impact of the dissociation from the cell culture surface and allow for the analysis of cellular functions in different phases of the growth curve (logarithmic phase vs. contact-inhibited phase7). Unfortunately, using microcarrier beads can be problematic due to the high stirring speed required in the respirometer and the potential for grinding of the beads along with cells underneath the stir bar. Despite these limitations, a broad range of applications remains compatible with calorespirometry. In particular, this method is poised to better facilitate pharmaceutical development by quickly identifying mitotoxic compounds in cultured cells.

In biological systems, the heat-release or enthalpy change during metabolism is typically monitored either directly (via direct calorimetry) or indirectly via O2 consumption and/or CO2 production (via respirometry). Unfortunately, when these techniques are used in isolation, critical information is lost, such as the contribution of anaerobic pathways to cellular metabolism. Calorespirometry is a powerful technique that relies on the concurrent measurement of both heat dissipation and respiration. Pioneering calorespirometric work investigated the anaerobic metabolism in fully oxygenated mammalian cells and demonstrated simultaneous contributions of both aerobic and anaerobic pathways to energy homeostasis despite the transformed cells being in a fully oxygenated environment1. Calorespirometry has since been applied to a wide variety of biological questions. Some examples include the study of animal energetics at low oxygen levels, the effects of both herbicide and estrogen on the gills of bivalves, the metabolism of terrestrial organisms, and the microbial decomposition of organic soil matter2,3,4,5,6. Furthermore, calorespirometry has revealed how metabolic preconditioning prior to freezing improves the cryopreservation of mammalian cells7. Each approach, both calorimetry and respirometry, has independently increased our knowledge of cellular and organismal bioenergetics. However, fundamental biological questions that can be answered through the use of calorespirometry remain relatively unexplored.
Hess&#39;s law states that the total enthalpy change of a reaction is independent of the pathway between the initial and final states. For example, the total enthalpy change for a biochemical pathway is the summation of the change in enthalpies of all reactions within the pathway. Calorimetry offers a real-time approach for measuring cellular heat production, which indiscriminately detects both aerobic and anaerobic pathways. This&#160;is based on the foundation that no energy is exchanged in the system except through the walls of the experimental ampule8. A change in heat dissipation is equal to the change in enthalpy released from all metabolic reactions in the ampule. Thus, a negative enthalpy correlates to a loss of heat from the system. Exhaustive research over the last four decades has characterized the thermodynamic landscape of both catabolism and anabolism. This is represented by a steady rise in research articles found under the search terms &#34;biological&#34; and &#34;calorimetry&#34; as indexed by the United States National Library of Medicine (NLM) at the National Institutes of Health (PubMed). The search reveals that prior to 1970, a total of 27 publications reference biological calorimetry; meanwhile, in 2016 alone, 546 publications utilized the technique.
Calorimetric methods are well established to determine heat production. However, more flexibility is granted for resolving the respirometric value. The respirometric measurement can consist of O2, CO2, or both O2 and CO2. Further, the measurement of O2 or CO2 can be accomplished by various techniques, including optrodes, Clark-type electrodes, and tunable diode laser absorption spectroscopy7,9,10,11. While CO2 production is a&#160;valuable metric in many respirometric studies, the medium for cultured cells often utilizes a bicarbonic buffer system for pH control12,13. To avoid complications of CO2 measurement in the bicarbonate system, the following protocol for the calorespirometry of cells in culture utilizes O2 as the sole respirometric parameter.
Concurrent with measuring oxygen flux, certain respirometers (see Table of Materials) are designed for detailed assessments of mitochondrial function. Substrate-uncoupler-inhibitor-titrations (SUIT) protocols are well established and are compatible with experiments designed to measure membrane potential or reactive oxygen species (ROS) formation14. The presented protocol for calorespirometry of intact cells is compatible with the introduction of chemical uncouplers such as carbonyl cyanide-p-trifluoromethoxyphenylhydrazone (FCCP) and the F0F1-ATP synthase inhibitor oligomycin. Through the addition of FCCP, oxygen consumption can be uncoupled from ATP production, which is useful to assess&#160;the impact of potential therapeutics on mitochondrial peformance15. Furthermore, the addition of oligomycin illuminates the extent of leak respiration. Thus, the respirometric measurements performed during calorespirometry are compatible with extensive protocols designed to further elucidate mitochondrial physiology.
The simultaneous measurement of both heat dissipation and oxygen flux allows for the calculation of the calorespirometric (CR) ratio. This ratio is then compared to the Thornton&#39;s constant or the theoretical oxycaloric equivalent, which ranges between -430 to -480 kJ mol-1 depending on the cell line or tissue of interest and the supplemented carbon substrates1,16. Thus, a more negative CR ratio reveals increased contributions from anaerobic pathways to overall metabolic activity. For example, the CR ratio for routine muscle tissue respiration without the active performance of work ranges from -448 to -468 kJ mol-1 which is within the range of the theoretical oxycaloric equivalent17,18. Meanwhile, mammalian cancer cells cultured in medium that is high in glucose display enhanced lactic acid fermentation following glycolysis and relatively low mitochondrial engagement19. This phenotype results in CR ratios in the range of -490 to -800 kJ mol-1, demonstrating a heightened involvement of the anaerobic pathways in the cellular metabolism as indicated by more negative CR ratios1,7,16,20.
Both commercial and non-profit cell and tissue distributors currently offer cell lines from over 150 species, with nearly 4,000 cell lines derived from humans. Immortalized cell lines are convenient tools for quickly evaluating the toxicity of potential therapeutics, many of which may directly or indirectly interfere with mitochondrial functions. Using transformed cells during drug screening may be of limited predictive value in part because of the Warburg effect, a hallmark of many cancers. Often, cancers generate ATP from substrate-level phosphorylation and maintain redox balance through the production of lactate without fully engaging the mitochondrion under aerobic conditions19. Pharmaceutical development is notoriously costly and inefficient, with approximately 8 out of 9 compounds tested in human clinical trials failing to achieve market approval21. While potential therapeutics may pass initial screening due to low cytotoxicity in cell lines, it is possible that some of these compounds are mitotoxic. Without a suitable method to detect how these toxins can impair the energy balance in primary cells that do not display the Warburg effect, critical information is often over-looked, bottlenecking therapeutic development in early stages.
Calorespirometry is a practical, noninvasive approach to analyze metabolic activity in a variety of biological samples, including cells and tissues. The core of the presented protocol is compatible with a range of applications. One complication, however, has been identified. Immortalized cells are often cultured in a glucose-free medium supplemented with galactose to increase the contribution of oxidative phosphorylation (OXPHOS) for energy production, in order to sensitize the cells to potential mitotoxins22,23. This metabolic reprogramming appears to obscure analysis when samples are placed in the stainless steel ampules used by the calorimeter15. Cells cultured in a glucose medium continue to engage in high metabolic activity for several hours. Meanwhile, cells cultured in galactose medium decrease the heat production within 30 min of their placement in the ampule, making measurements restricted to early experimental time points. This behavior, unfortunately, hinders the opportunity to assess their cellular proliferation. Despite this specific limitation, most applications are compatible with calorespirometric analysis and detailed metabolic information can be obtained through this approach.

<table border="0" cellpadding="0" cellspacing="0" width="627">
	<tbody>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">HepG2 Cells</td>
			<td style="width:125px;">American Type Culture Collection</td>
			<td style="width:119px;">HB-8065</td>
			<td style="width:167px;">Cells used for calorespirometry</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">O2k-Respirometer</td>
			<td style="width:125px;">Oroboros Instruments</td>
			<td style="width:119px;">10022-02</td>
			<td>Respirometer</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">LKB 2277 thermal activity monitor (TAM)</td>
			<td style="width:125px;">Thermometric AB</td>
			<td style="width:119px;"></td>
			<td>Thermometric was purchased by TA Instruments</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Sodium Pyruvate (100 mM)</td>
			<td style="width:125px;">Thermofisher Scientific</td>
			<td style="width:119px;">11360070</td>
			<td>100x solution added to DMEM medium</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Fetal Bovine Serum - Premiuim Select</td>
			<td style="width:125px;">Atlanta Biologicals</td>
			<td style="width:119px;">S11550</td>
			<td>Added to 10% in DMEM medium</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Trypsn-EDTA (0.25%)</td>
			<td style="width:125px;">Thermofisher Scientific</td>
			<td style="width:119px;">25200072</td>
			<td>Cell dissociation reagent</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Oligomycin from Streptomyces diastatochromogenes</td>
			<td style="width:125px;">Sigma Aldrich&#160;</td>
			<td style="width:119px;">O4876</td>
			<td>Mitochondrial Inhibitor</td>
		</tr>
		<tr height="63">
			<td height="63" style="height:63px;width:216px;">Carbonyl cyanide 4-(trifluoromethoxy)phenylhydrazone</td>
			<td style="width:125px;">Sigma Aldrich</td>
			<td style="width:119px;">C2920</td>
			<td>Mitochondrial Uncoupler</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Corning 100 mm TC-Treated Culture Dish</td>
			<td style="width:125px;">Corning Corporation</td>
			<td style="width:119px;">430167</td>
			<td>Tissue culture dish</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Glucose, powder</td>
			<td style="width:125px;">Thermofisher Scientific</td>
			<td style="width:119px;">15023021</td>
			<td>Glucose for DMEM medium</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Galactose, powder</td>
			<td style="width:125px;">Fischer Scientific</td>
			<td style="width:119px;">BP656500</td>
			<td>Galactose for DMEM medium</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">L-Glutamine (200 mM)</td>
			<td style="width:125px;">Thermofisher Scientific</td>
			<td style="width:119px;">25030081</td>
			<td>Glutamine for DMEM medium</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">DMEM, no glucose</td>
			<td style="width:125px;">Thermofisher Scientific</td>
			<td style="width:119px;">11966025</td>
			<td>Cell culture medium</td>
		</tr>
	</tbody>
</table>

calorespirometry: a powerful, noninvasive approach to investigate cellular energy metabolism

Many cell lines used in basic biological and biomedical research maintain energy homeostasis through a combination of both aerobic and anaerobic respiration. However, the extent to which both pathways contribute to the landscape of cellular energy production is consistently overlooked. Transformed cells cultured in saturating levels of glucose often show a decreased dependency on oxidative phosphorylation for ATP production, which is compensated by an increase in substrate-level phosphorylation. This shift in metabolic poise allows cells to proliferate despite the presence of mitochondrial toxins. In neglecting the altered metabolic poise of transformed cells, results from a pharmaceutical screening may be misinterpreted since the potentially mitotoxic effects may not be detected using model cell lines cultured in the presence of high glucose concentrations. This protocol describes the pairing of two powerful techniques, respirometry and calorimetry, which allows for the quantitative and noninvasive assessment of both aerobic and anaerobic contributions to cellular ATP production. Both aerobic and anaerobic respirations generate heat, which can be monitored via calorimetry. Meanwhile, measuring the rate of oxygen consumption can assess the extent of aerobic respiration. When both heat dissipation and oxygen consumption are measured simultaneously, the calorespirometric ratio can be determined. The experimentally obtained value can then be compared to the theoretical oxycaloric equivalent and the extent of the anaerobic respiration can be judged. Thus, calorespirometry provides a unique method to analyze a wide range of biological questions, including drug development, microbial growth, and fundamental bioenergetics under both normoxic and hypoxic conditions.

The reproducibility of calorespirometric measurements depends on a proper and consistent sample preparation. Samples prepared from cell culture should not be used if the plates are overgrown, as cell counts can become inaccurate due to clumping. Further, reduced heat flows due to limited substrate diffusion in the clumped cells may occur. Therefore, when using adherent cells, it is critical to select a plate with the confluency between 60 - 80% and to change the medium 24 h prior to the experiment.
Proper instrumental calibration and maintenance are also critical for informative and reproducible calorespirometric measurements. By following the manufacturer&#39;s established cleaning protocol for the respirometer, extensive washes with ethanol are implemented to ensure that residual hydrophobic uncouplers and inhibitors are removed and do not compromise the measurements by partially inhibiting or uncoupling mitochondria.&#160;When the respirometer chamber is operating prior to the sample measurement, both the oxygen concentration and the oxygen flux traces should be stable as shown in Figure 1. If the POS is in need of sensor service, the oxygen flux will not stabilize as seen in Figure 2.&#160;Measurements should not be performed when the POS is not properly serviced. Additionally, the ampules utilized in the calorimeter should be cleaned, sterilized, and dried prior to use, as biological contamination can interfere with the heat production of the sample.
Cell samples are prepared after the proper calibration and preparation of both the respirometer and the calorimeter. First, the stopper in the respirometer is closed and the chamber is inspected to ensure no air bubbles are present. Then, HepG2 cells are concentrated to ~2 x 106/50 &#181;L for respirometry and immediately placed on ice. For calorimetry, HepG2 cells from the concentrated sample are diluted in equilibrated DMEM at&#160;~1 x 105 cells/mL. After cells are thoroughly mixed by pipetting, 2.5 mL of the suspension is added to a sterile ampule. The ampule is tightly sealed, and any external debris is removed by blowing the ampule with a stream of air from an air pump. The ampule is then slowly lowered to position 1, where it remains for 15 min. After 15 min has passed, both the reference and sample ampules are then slowly lowered to position 2, and measurements can begin to be recorded once the heat output is stable (Figure 3).
A final cell count is performed after the cells are added to both the calorimeter and the respirometer to determine the precise number of cells introduced into the calorimeter. The heat production is then expressed per million cells. If the HepG2 cells are cultured in the absence of glucose and galactose is supplemented to increase&#160;mitochondrial involvement, the cells do not proliferate inside the calorimeter, but instead decrease their metabolic activity after about 30 min in the ampule. This restricts the CR calculation to the earliest time point at which the heat dissipation was measured (Figure 4).
It is critical to record the time the cells are added to the ampule so that the measurements of heat production and oxygen consumption are collected at identical time points. If this precaution is not taken, important experimental errors can be introduced in the determination of the CR ratio. During the 15-min period, while the ampule is at position 1 in the calorimeter, the cells are sufficiently mixed by pipetting them, and 2 x 106 cells are injected into the respirometer via a Hamilton syringe. Oxygen flux stabilizes after approximately 15 min and oxygen concentration steadily declines (Figure 5). Again, it is critical that the time of the sample injection is recorded for data analysis. The stabilized oxygen flux of the cells serves as the surrogate for oxygen consumption when calculating the CR ratio. Additional titrations are performed to assess the leak respiration (1 &#181;L oligomycin) indicated by a drop to a lower oxygen flux and the maximal uncoupled respiration (3 - 5 titrations of 2 &#181;L FCCP) indicated by the failure to increase the respiration after successive FCCP titrations. An incomplete uncoupling or an inhibition of respiration may not be observed if the FCCP stock has expired (Figure 6).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/57724/57724fig1.jpg" /> 
Figure 1: 100% air calibration of respirometer and status of polarographic oxygen sensor (POS). The signals for the oxygen concentration (blue line) and the oxygen flux (red line) both stabilize prior to the 100% air calibration. The stable lines indicate that the respirometer is ready for calibration. The calibration should be performed in DMEM, and not in H2O. <a href="/files/ftp_upload/57724/57724fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/57724/57724fig2.jpg" /> 
Figure 2: POS in need of service. After the stabilization of the oxygen concentration (blue trace), the oxygen flux (red trace) fails to stabilize. The POS should not be used for experimentation and service should be applied as described by the manufacturer. An increased instability of the oxygen flux correlates with an increased demand for POS service. <a href="/files/ftp_upload/57724/57724fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/57724/57724fig3v2.jpg" /> 
Figure 3: Approximately 2.5 x 105 HepG2 cells cultured in glucose medium show signs of proliferation in the calorimeter. Loading ~2.5 x 105 HepG2 cells cultured in the presence of glucose per ampule generates a constant heat production of -20 to -28 &#181;W using the 30-&#181;W setting on the instrument and an increased heat production over time correlates with cellular proliferation inside the ampule. <a href="/files/ftp_upload/57724/57724fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/57724/57724fig4v2.jpg" /> 
Figure 4: Approximately 2.5 x 105 HepG2 cells cultured in galactose medium do not proliferate and heat production decreases over time. Loading ~2.5 x 105 HepG2 cells cultured in galactose per ampule results in an immediate heat production of ~-20 &#181;W. However, the heat production decreases over time and restricts CR-ratio measurements to the early time points of the experiment. <a href="/files/ftp_upload/57724/57724fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/57724/57724fig5.jpg" /> 
Figure 5: Routine, uncoupled, and leak respiration of HepG2 cells. Once cells were injected into the respirometer, oxygen concentration steadily decreased (blue trace). After the stabilization of the oxygen flux (red trace) during the routine respiration of intact cells, oligomycin was added to inhibit the F0F1-ATP synthase. This is indicated by the stabilization of&#160;oxygen flux after oligomycin addition, which reveals leak respiration. Approximately 3 - 5 titrations of FCCP should be performed to uncouple&#160;respiration from&#160;ATP synthesis&#160;and to reveal&#160;electron transport system (ETS) capacity. <a href="/files/ftp_upload/57724/57724fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/57724/57724fig6.jpg" /> 
Figure 6: Unsuccessful uncoupling of HepG2 cell respiration. After addition of cells, oxygen flux stabilized (red trace) during routine respiration and oxygen concentration steadily decreased (blue line). Titration of FCCP resulted in an incomplete uncoupling and eventual inhibition, preventing the maximal ETS capacity from being reached. FCCP was likely contaminated or degraded from long-term storage. If observed, a fresh FCCP stock must be prepared. Also, confirm that&#160;oligomycin does not inhibit&#160;ETS capacity during uncoupling. <a href="/files/ftp_upload/57724/57724fig6large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Calorespirometry: A Powerful, Noninvasive Approach to Investigate Cellular Energy Metabolism at JoVE.com

Calorespirometry: A Powerful, Noninvasive Approach to Investigate Cellular Energy Metabolism

This protocol describes calorespirometry, the direct and simultaneous measurement of both heat dissipation and respiration, which provides a noninvasive approach to assess energy metabolism. This technique is used to assess the contribution of both aerobic and anaerobic pathways to energy utilization by monitoring the total cellular energy flow.

Many cell lines used in basic biological and biomedical research maintain energy homeostasis through a combination of both aerobic and anaerobic ...

calorespirometry-powerful-noninvasive-approach-to-investigate

Nanyang Technological University

Research

JoVE Journal

Bioengineering

7.5K Views.  University of Louisville. This protocol describes calorespirometry, the direct and simultaneous measurement of both heat dissipation and respiration, which provides a noninvasive approach to assess energy metabolism. This technique is used to assess the contribution of both aerobic and anaerobic pathways to energy utilization by monitoring the total cellular energy flow.

Video: Calorespirometry: A Powerful, Noninvasive Approach to Investigate Cellular Energy Metabolism

The Three-Dimensional Human Skin Reconstruct Model: a Tool to Study Normal  Skin and Melanoma Progression

Most in vitro studies in experimental skin biology have been done in 2-dimensional (2D) monocultures, while accumulating evidence suggests that cells behave differently when they are grown within a 3D extra-cellular matrix and also interact with other cells (1-5). Mouse models have been broadly utilized to study tissue morphogenesis in vivo. However mouse and human skin have significant differences in cellular architecture and physiology, which makes it difficult to extrapolate mouse studies to humans. Since melanocytes in mouse skin are mostly localized in hair follicles, they have distinct biological properties from those of humans, which locate primarily at the basal layer of the epidermis. The recent development of 3D human skin reconstruct models has enabled the field to investigate cell-matrix and cell-cell interactions between different cell types. The reconstructs consist of a "dermis" with fibroblasts embedded in a collagen I matrix, an "epidermis", which is comprised of stratified, differentiated keratinocytes and a functional basement membrane, which separates epidermis from dermis. Collagen provides scaffolding, nutrient delivery, and potential for cell-to-cell interaction. The 3D skin models incorporating melanocytic cells recapitulate natural features of melanocyte homeostasis and melanoma progression in human skin. As in vivo, melanocytes in reconstructed skin are localized at the basement membrane interspersed with basal layer keratinocytes. Melanoma cells exhibit the same characteristics reflecting the original tumor stage (RGP, VGP and metastatic melanoma cells) in vivo. Recently, dermal stem cells have been identified in the human dermis (6). These multi-potent stem cells can migrate to the epidermis and differentiate to melanocytes.

In this report, we describe the three-dimensional skin reconstruct model which mimics human skin in architecture and composition. Melanocyte physiology, melanoma progression and the fate of dermal stem cells have been investigated using the skin reconstruct model. The model is also useful as a preclinical tool for drug assessment.

Most in vitro studies in experimental skin biology have been done in 2-dimensional (2D) monocultures, while accumulating evidence suggests that cells ...

Design of a Cyclic Pressure Bioreactor for the Ex Vivo Study of Aortic Heart Valves

The aortic valve, located between the left ventricle and the aorta, allows for unidirectional blood flow, preventing backflow into the ventricle. Aortic valve leaflets are composed of interstitial cells suspended within an extracellular matrix (ECM) and are lined with an endothelial cell monolayer. The valve withstands a harsh, dynamic environment and is constantly exposed to shear, flexion, tension, and compression. Research has shown calcific lesions in diseased valves occur in areas of high mechanical stress as a result of endothelial disruption or interstitial matrix damage1-3. Hence, it is not surprising that epidemiological studies have shown high blood pressure to be a leading risk factor in the onset of aortic valve disease4. 
The only treatment option currently available for valve disease is surgical replacement of the diseased valve with a bioprosthetic or mechanical valve5. Improved understanding of valve biology in response to physical stresses would help elucidate the mechanisms of valve pathogenesis. In turn, this could help in the development of non-invasive therapies such as pharmaceutical intervention or prevention. Several bioreactors have been previously developed to study the mechanobiology of native or engineered heart valves6-9. Pulsatile bioreactors have also been developed to study a range of tissues including cartilage10, bone11 and bladder12. The aim of this work was to develop a cyclic pressure system that could be used to elucidate the biological response of aortic valve leaflets to increased pressure loads. 
The system consisted of an acrylic chamber in which to place samples and produce cyclic pressure, viton diaphragm solenoid valves to control the timing of the pressure cycle, and a computer to control electrical devices. The pressure was monitored using a pressure transducer, and the signal was conditioned using a load cell conditioner. A LabVIEW program regulated the pressure using an analog device to pump compressed air into the system at the appropriate rate. The system mimicked the dynamic transvalvular pressure levels associated with the aortic valve; a saw tooth wave produced a gradual increase in pressure, typical of the transvalvular pressure gradient that is present across the valve during diastole, followed by a sharp pressure drop depicting valve opening in systole. The LabVIEW program allowed users to control the magnitude and frequency of cyclic pressure. The system was able to subject tissue samples to physiological and pathological pressure conditions. This device can be used to increase our understanding of how heart valves respond to changes in the local mechanical environment.

Design of a Cyclic Pressure Bioreactor for the Ex Vivo Study of Aortic Heart Valves

A cyclic pressure bioreactor capable of subjecting heart valve tissue to physiological and pathological pressure conditions has been designed. A LabVIEW program allows users to control pressure magnitude, amplitude and frequency. This device can be used to study the mechanobiology of heart valve tissue or isolated cells.

The aortic valve, located between the left ventricle and the aorta, allows for unidirectional blood flow, preventing backflow into the ventricle. Aortic ...

Directed Cellular Self-Assembly to Fabricate Cell-Derived Tissue Rings for Biomechanical Analysis and Tissue Engineering

Each year, hundreds of thousands of patients undergo coronary artery bypass surgery in the United States.1 Approximately one third of these patients do not have suitable autologous donor vessels due to disease progression or previous harvest. The aim of vascular tissue engineering is to develop a suitable alternative source for these bypass grafts. In addition, engineered vascular tissue may prove valuable as living vascular models to study cardiovascular diseases. Several promising approaches to engineering blood vessels have been explored, with many recent studies focusing on development and analysis of cell-based methods.2-5 Herein, we present a method to rapidly self-assemble cells into 3D tissue rings that can be used in vitro to model vascular tissues.
To do this, suspensions of smooth muscle cells are seeded into round-bottomed annular agarose wells. The non-adhesive properties of the agarose allow the cells to settle, aggregate and contract around a post at the center of the well to form a cohesive tissue ring.6,7 These rings can be cultured for several days prior to harvesting for mechanical, physiological, biochemical, or histological analysis. We have shown that these cell-derived tissue rings yield at 100-500 kPa ultimate tensile strength8 which exceeds the value reported for other tissue engineered vascular constructs cultured for similar durations (&lt;30 kPa).9,10 Our results demonstrate that robust cell-derived vascular tissue ring generation can be achieved within a short time period, and offers the opportunity for direct and quantitative assessment of the contributions of cells and cell-derived matrix (CDM) to vascular tissue structure and function.

This article outlines a versatile method to create cell-derived tissue rings by cellular self-assembly. Smooth muscle cells seeded into ring-shaped agarose wells aggregate and contract to form robust three-dimensional (3D) tissues within 7 days. Millimeter-scale tissue rings are conducive to mechanical testing and serve as building blocks for tissue assembly.

Each year, hundreds of thousands of patients undergo coronary artery bypass surgery in the United States.1 Approximately one third of these patients do ...

FIBS-enabled Noninvasive Metabolic Profiling

In the era of computational biology, new high throughput experimental systems are necessary in order to populate and refine models so that they can be validated for predictive purposes. Ideally such systems would be low volume, which precludes sampling and destructive analyses when time course data are to be obtained. What is needed is an in situ monitoring tool which can report the necessary information in real-time and noninvasively. An interesting option is the use of fluorescent, protein-based in vivo biological sensors as reporters of intracellular concentrations. One particular class of in vivo biosensors that has found applications in metabolite quantification is based on F&#246;rster Resonance Energy Transfer (FRET) between two fluorescent proteins connected by a ligand binding domain. FRET integrated biological sensors (FIBS) are constitutively produced within the cell line, they have fast response times and their spectral characteristics change based on the concentration of metabolite within the cell. In this paper, the method for constructing Chinese hamster ovary (CHO) cell lines that constitutively express a FIBS for glucose and glutamine and calibrating the FIBS in vivo in batch cell culture in order to enable future quantification of intracellular metabolite concentration is described. Data from fed-batch CHO cell cultures demonstrates that the FIBS was able in each case to detect the resulting change in the intracellular concentration. Using the fluorescent signal from the FIBS and the previously constructed calibration curve, the intracellular concentration was accurately determined as confirmed by an independent enzymatic assay.

A description of how to calibrate F&#246;rster Resonance Energy Transfer integrated biological sensors (FIBS) for in situ metabolic profiling is presented.&#160;The FIBS can be used to measure intracellular levels of metabolites noninvasively aiding in the development of metabolic models and high throughput screening of bioprocess conditions.

In the era of computational biology, new high throughput experimental systems are necessary in order to populate and refine models so that they can be ...

From Voxels to Knowledge: A Practical Guide to the Segmentation of Complex Electron Microscopy 3D-Data

Modern 3D electron microscopy approaches have recently allowed unprecedented insight into the 3D ultrastructural organization of cells and tissues, enabling the visualization of large macromolecular machines, such as adhesion complexes, as well as higher-order structures, such as the cytoskeleton and cellular organelles in their respective cell and tissue context. Given the inherent complexity of cellular volumes, it is essential to first extract the features of interest in order to allow visualization, quantification, and therefore comprehension of their 3D organization. Each data set is defined by distinct characteristics, e.g., signal-to-noise ratio, crispness (sharpness) of the data, heterogeneity of its features, crowdedness of features, presence or absence of characteristic shapes that allow for easy identification, and the percentage of the entire volume that a specific region of interest occupies. All these characteristics need to be considered when deciding on which approach to take for segmentation.
The six different 3D ultrastructural data sets presented were obtained by three different imaging approaches: resin embedded stained electron tomography, focused ion beam- and serial block face- scanning electron microscopy (FIB-SEM, SBF-SEM) of mildly stained and heavily stained samples, respectively. For these data sets, four different segmentation approaches have been applied: (1) fully manual model building followed solely by visualization of the model, (2) manual tracing segmentation of the data followed by surface rendering, (3) semi-automated approaches followed by surface rendering, or (4) automated custom-designed segmentation algorithms followed by surface rendering and quantitative analysis. Depending on the combination of data set characteristics, it was found that typically one of these four categorical approaches outperforms the others, but depending on the exact sequence of criteria, more than one approach may be successful. Based on these data, we propose a triage scheme that categorizes both objective data set characteristics and subjective personal criteria for the analysis of the different data sets.

The bottleneck for cellular 3D electron microscopy is feature extraction (segmentation) in highly complex 3D density maps. We have developed a set of criteria, which provides guidance regarding which segmentation approach (manual, semi-automated, or automated) is best suited for different data types, thus providing a starting point for effective segmentation.

Modern 3D electron microscopy approaches have recently allowed unprecedented insight into the 3D ultrastructural organization of cells and tissues, ...

Planar Gradient Diffusion System to Investigate Chemotaxis in a 3D Collagen Matrix

The importance of cell migration can be seen through the development of human life. When cells migrate, they generate forces and transfer these forces to their surrounding area, leading to cell movement and migration. In order to understand the mechanisms that can alter and/or affect cell migration, one can study these forces. In theory, understanding the fundamental mechanisms and forces underlying cell migration holds the promise of effective approaches for treating diseases and promoting cellular transplantation. Unfortunately, modern chemotaxis chambers that have been developed are usually restricted to two dimensions (2D) and have complex diffusion gradients that make the experiment difficult to interpret. To this end, we have developed, and describe in this paper, a direct-viewing chamber for chemotaxis studies, which allows one to overcome modern chemotaxis chamber obstacles able to measure cell forces and specific concentration within the chamber in a 3D environment to study cell 3D migration. More compelling, this approach allows one to successfully model diffusion through 3D collagen matrices and calculate the coefficient of diffusion of a chemoattractant through multiple different concentrations of collagen, while keeping the system simple and user friendly for traction force microscopy (TFM) and digital volume correlation (DVC) analysis.

Cell migration is an important part of human development and life. In order to understand the mechanisms that can alter cell migration, we present a planar gradient diffusion system to investigate chemotaxis in a 3D collagen matrix, which allows one to overcome modern diffusion chamber limitations of existing assays.

The importance of cell migration can be seen through the development of human life. When cells migrate, they generate forces and transfer these forces to ...

An Experimental and Finite Element Protocol to Investigate the Transport of Neutral and Charged Solutes across Articular Cartilage

Osteoarthritis (OA) is a debilitating disease that is associated with degeneration of articular cartilage and subchondral bone. Degeneration of articular cartilage impairs its load-bearing function substantially as it experiences tremendous chemical degradation, i.e. proteoglycan loss and collagen fibril disruption. One promising way to investigate chemical damage mechanisms during OA is to expose the cartilage specimens to an external solute and monitor the diffusion of the molecules. The degree of cartilage damage (i.e. concentration and configuration of essential macromolecules) is associated with collisional energy loss of external solutes while moving across articular cartilage creates different diffusion characteristics compared to healthy cartilage. In this study, we introduce a protocol, which consists of several steps and is based on previously developed experimental micro-Computed Tomography (micro-CT) and finite element modeling. The transport of charged and uncharged iodinated molecules is first recorded using micro-CT, which is followed by applying biphasic-solute and multiphasic finite element models to obtain diffusion coefficients and fixed charge densities across cartilage zones.

We propose a protocol to investigate the transport of charged and uncharged molecules across articular cartilage with the aid of recently developed experimental and numerical methods.

Osteoarthritis (OA) is a debilitating disease that is associated with degeneration of articular cartilage and subchondral bone. Degeneration of articular ...

Workflow Based on the Combination of Isotopic Tracer Experiments to Investigate Microbial Metabolism of Multiple Nutrient Sources

Studies in the field of microbiology rely on the implementation of a wide range of methodologies. In particular, the development of appropriate methods substantially contributes to providing extensive knowledge of the metabolism of microorganisms growing in chemically defined media containing unique nitrogen and carbon sources. In contrast, the management through metabolism of multiple nutrient sources, despite their broad presence in natural or industrial environments, remains virtually unexplored. This situation is mainly due to the lack of suitable methodologies, which hinders investigations.
We report an experimental strategy to quantitatively and comprehensively explore how metabolism operates when a nutrient is provided as a mixture of different molecules, i.e., a complex resource. Here, we describe its application for assessing the partitioning of multiple nitrogen sources through the yeast metabolic network. The workflow combines information obtained during stable isotope tracer experiments using selected 13C- or 15N-labeled substrates. It first consists of parallel and reproducible fermentations in the same medium, which includes a mixture of N-containing molecules; however,a selected nitrogen source is labeled each time. A combination of analytical procedures (HPLC, GC-MS) is implemented to assess the labeling patterns of targeted compounds and to quantify the consumption and recovery of substrates in other metabolites. An integrated analysis of the complete dataset provides an overview of the fate of consumed substrates within cells. This approach requires an accurate protocol for the collection of samples&#8211;facilitated by a robot-assisted system for online monitoring of fermentations&#8211;and the achievement of numerous time-consuming analyses. Despite these constraints, it allowed understanding, for the first time, the partitioning of multiple nitrogen sources throughout the yeast metabolic network. We elucidated the redistribution of nitrogen from more abundant sources toward other N-compounds and determined the metabolic origins of volatile molecules and proteinogenic amino acids.

This protocol describes an experimental procedure to quantitatively and comprehensively investigate the metabolism of multiple nutrient sources. This workflow, based on a combination of isotopic tracer experiments and an analytical procedure, allows the fate of consumed nutrients and the metabolic origin of molecules synthetized by microorganisms to be determined.

Studies in the field of microbiology rely on the implementation of a wide range of methodologies. In particular, the development of appropriate methods ...

Fabrication of a Multiplexed Artificial Cellular MicroEnvironment Array

Cellular microenvironments consist of a variety of cues, such as growth factors, extracellular matrices, and intercellular interactions. These cues are well orchestrated and are crucial in regulating cell functions in a living system. Although a number of researchers have attempted to investigate the correlation between environmental factors and desired cellular functions, much remains unknown. This is largely due to the lack of a proper methodology to mimic such environmental cues in vitro, and simultaneously test different environmental cues on cells. Here, we report an integrated platform of microfluidic channels and a nanofiber array, followed by high-content single-cell analysis, to examine stem cell phenotypes altered by distinct environmental factors. To demonstrate the application of this platform, this study focuses on the phenotypes of self-renewing human pluripotent stem cells (hPSCs). Here, we present the preparation procedures for a nanofiber array and the microfluidic structure in the fabrication of a Multiplexed Artificial Cellular MicroEnvironment (MACME) array. Moreover, overall steps of the single-cell profiling, cell staining with multiple fluorescent markers, multiple fluorescence imaging, and statistical analyses, are described.

This article describes the detailed methodology to prepare a Multiplexed Artificial Cellular MicroEnvironment (MACME) array for high-throughput manipulation of physical and chemical cues mimicking in vivo cellular microenvironments and to identify the optimal cellular environment for human pluripotent stem cells (hPSCs) with single-cell profiling.

Cellular microenvironments consist of a variety of cues, such as growth factors, extracellular matrices, and intercellular interactions. These cues are ...

Nanosensors to Detect Protease Activity In Vivo for Noninvasive Diagnostics

Proteases are multi-functional enzymes that specialize in the hydrolysis of peptide-bonds and control broad biological processes including homeostasis and allostasis. Moreover, dysregulated protease activity drives pathogenesis and is a functional biomarker of diseases such as cancer; therefore, the ability to detect protease activity in vivo may provide clinically relevant information for biomedical diagnostics. The goal of this protocol is to create nanosensors that probe for protease activity in vivo by producing a quantifiable signal in urine. These protease nanosensors consist of two components: a nanoparticle and substrate. The nanoparticle functions to increase circulation half-life and substrate delivery to target disease sites. The substrate is a short peptide sequence (6-8 AA), which is designed to be specific to a target protease or group of proteases. The substrate is conjugated to the surface of the nanoparticle and is terminated by a reporter, such as a fluorescent marker, for detection. As dysregulated proteases cleave the peptide substrate, the reporter is filtered into urine for quantification as a biomarker of protease activity. Herein we describe construction of a nanosensor for matrix metalloproteinase 9 (MMP9), which is associated with tumor progression and metastasis, for detection of colorectal cancer in a mouse model.

Nanosensors to Detect Protease Activity In Vivo for Noninvasive Diagnostics

Proteases are tightly regulated enzymes involved in fundamental biological processes, and dysregulated protease activity drives progression of complex diseases such as cancer. This method&#39;s goal is to create nanosensors that measure protease activity in vivo by producing a cleavage signal that is detectable from host urine and discriminates disease.

Proteases are multi-functional enzymes that specialize in the hydrolysis of peptide-bonds and control broad biological processes including homeostasis and ...