We thank Andreas Ladurner, Carla Margulis and their teams for extensive experimental support. We thank Caitlin Ondracek for her comments on the manuscript. We would like to thank Sofia Vikstrom for assisting us in establishing the early phases of this technique. We also thank May Sanderhoff for her technical help. LB is funded by the German Federal Ministry of Education and Research (Infrafrontier grant 01KX1012). SP was funded by an AXA Research Fund postdoctoral fellowship and the NSFC (Grant number 81870900). AVV is funded by the QBM.

1. Instrument Preparation
Note: For this experiment, we have used a Seahorse XF24 device with &#8220;islet plates&#8221;. The operation of the technique uses different cycles of mixing, waiting and measurements as well as the possibility to add substances to the measurement compartment.
<ol>
	<li>Turn on the machine well before the start of the experiment so that there is adequate time to reach the desired temperature and remain stable.</li>
	<li>In the software setup (administration mode), choose the length of the cartridge calibration (here, 20 min was chosen) and the desired temperature. 
	Note: While mitochondria or mammalian tissue measurements are typically carried out at 37 &#176;C, fly head ambient temperature is 25 &#176;C but results of measurements at 31 &#176;C are published. We used 31 &#176;C since this is the lowest temperature setting for the device at room temperature. To reach temperatures of 25 &#176;C or lower, place the machine in a colder room or at 11 &#176;C as recently published21.</li>
	<li>In the software, use the following protocol: 3 min mixing &#8211; 2 min waiting &#8211; 2 min measuring. Depending on the experimental design, add injection steps from ports A-D after a chosen measurement step.
	<ol>
		<li>For quality check, and determination of basal OCR, wait for at least three measurement cycles before injecting the first drug via port A. For a detailed timeline of the protocol, please see reference Becker et al.13.</li>
	</ol>
	</li>
</ol>
2. Cartridge Preparation
<ol>
	<li>Pre-calibrate the cartridge a day (or at least 4 h) prior to testing. Add 1.0 mL of Calibrant (pH 7.4) to each well and place the sensor cartridge on top of the plate and store at 37 &#176;C without CO2 for overnight or up to 72 h. Prevent the evaporation of cartridge with parafilm if it is being hydrated for more than 24 h.</li>
	<li>Ensure that the experimental drugs are well dissolved in the medium (fresh medium + 2.5% glucose) before the start of the experiment.</li>
	<li>Measure and adjust the pH of the drug solution to the pH of the vehicle at the desired temperature to avoid any pH difference during drug injection.</li>
	<li>Pipet the drug solution to its allocated injection port. For example, use 77 &#956;L for port A to achieve a 1:10 dilution in a 770 &#956;L solution and subsequently 85 &#956;L for port B.</li>
	<li>Load the cartridge into the machine and begin calibration.</li>
</ol>
3. Plate Preparation
Note: It is strongly recommended that two people prepare the plate simultaneously. The duration of one plate preparation per two people may require ~45-60 minutes.
<ol>
	<li>Adjust the freshly prepared media + 2.5% glucose to the desired pH with 1 N HCl. Make sure that the pH is not affected by changes in temperature.</li>
	<li>Prepare an ice box and place a metal plate on the ice.</li>
	<li>Open the islet plate (the 24-well plate) package and immerse the nets in a Petri dish (92 mm x 16 mm) with media.</li>
	<li>Collect one net with the inserter (a small instrument that places the net firmly in the well) and have the inserter stand up next to the microscope. Add a small drop of media to the net attached to the inserter.</li>
	<li>Anesthetize the flies (one week or 4 weeks old canton males were used here) by placing the flies on the ice-cold metal plate.</li>
	<li>Using forceps, grab the abdomen of a fly and immerse it in the media in a Petri dish under the microscope.</li>
	<li>Using a second pair of forceps, gently remove the head of the fly. Place it in the middle of the net attached to the inserter and verify that the head is immersed in media.</li>
	<li>Center the heads when there are 16 of them on the net. Remove superfluous fluid before centering the heads to prevent loss of heads while placing them in the well. 
	Note: 16 fly heads were used as this number gave sufficient and stable data within a reasonable time of plate preparation during method establishment.</li>
	<li>Using the inserter, place the net in the well. Ensure that the heads are trapped under the net. Slowly add 700 &#956;L of media + 2.5% glucose (Figure 1). Repeat the process for each of the wells. 
	Note: 20 wells of fly head samples and 4 empty wells for background calibration per plate is recommended. Make sure that empty wells also contain a net with 700 &#956;L of the buffer + 2.5% glucose.</li>
	<li>Check the wells for air bubbles under the nets via the microscope. Pipet gently up and down using a 1 mL pipet to remove any bubbles. Keep the heads centered for a reliable OCR reading.</li>
	<li>Add the plate to the machine and start the measurement.</li>
</ol>
4. Analysis of the OCR Measurements
<ol>
	<li>At the end of the protocol, remove the cartridge.</li>
	<li>As a quality check, observe any visible leftovers in the port fillings. Discard the cartridge and plate (Option 1) if the heads are not to be used for protein extraction, e.g., (see Option 2).</li>
	<li>Extract the spreadsheet files and quality check each well for oxygen and pH levels. Make sure that the background wells show no OCR and that oxygen levels are stable.
	<ol>
		<li>Use an algorithm for data analysis, some of which can be chosen in the respective software. Use the AKOS algorithm for OCR values2 if the range of oxygen levels during the entire measurement, between the first and last tick (= sub-measurement) are similar between two biological samples and the oxygen levels of the last ticks are not lower than 95 (mmHg) (the heads&#8217; OCR is markedly lower at this oxygen level), (Figure 2).</li>
		<li>Some conditions will cause the sample to generate a rapid OCR and may display lower oxygen levels during the 1st tick and/or in the last tick (anoxia) (Figure 3A). In such a scenario, use an alternate measurement method such as the FIXED algorithm. In anoxia, the OCR is greatly reduced due to low levels of oxygen in the solution. As such, the AKOS algorithm yields misleading readings. 
		Note: The newer machine lacks the FIXED algorithm. Therefore, it is preferred to extract the total oxygen levels and plot the rate per time for the first 3-5 ticks (Figure 3).</li>
	</ol>
	</li>
</ol>
5. (Option 2) Biochemical Analysis of the Head Segment
<ol>
	<li>To measure the biochemical (metabolites, proteome, etc.) properties of a head segment, adjust the run time to the requirement; however, it is possible to abort the protocol at any time and remove the plate.</li>
	<li>Once the plate is removed, use non-sharpened forceps to make a hole in the net and remove it thereby releasing the heads to float.</li>
	<li>Using a 1 mL pipet with a cut tip and transfer the heads to a vial.</li>
	<li>Quickly discard the buffer and snap-freeze the heads in liquid nitrogen. Store the heads at -80 &#176;C for future analysis.</li>
</ol>

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The authors declare that they have no competing financial interests.

Our new technique offers a novel approach to study metabolic changes in aging and disease in the context of whole fly head segments22. The method can also be suited to study the impact of KDAC sodium butyrate on oxygen consumption. As we have demonstrated, lysine deacetlyase inhibitors (HDACs/KDACs) result in OCR changes. Essentially, as the targets of such inhibitors are normally not localized in the mitochondria (these inhibitors do not impact the Class III deacetylases, the Sirtuins)24, such drugs could only be tested on an at least tissue level. Indeed, various drugs are injected directly to the brain, thus bypassing possible processing/modification/inactivation by the digestive system. As such, our technique offers novel insight into how such drugs directly impact the head segment.
There are several critical steps. First, as stated in the protocol, we highly recommend preparing a plate under one hour, with two pairs of hands preparing the plate. From our experience, the quality and stability of the OCR measurements are better when prepared in a timely manner. When taking too long, the occurrence of low OCR consuming wells is increasing, as well as shorter duration of stable OCR. Second, it is important to conduct a quality check and ensure that the experimental conditions between various samples are similar (pH, oxygen levels). Finally, a critical step is choosing the correct algorithm to analyze the samples. As we have demonstrated, the default AKOS algorithm yielded a misleading and sometimes opposing calculation in samples that consumed oxygen at high rates13. We therefore stress the importance of checking the raw data for oxygen levels and comparing the resulting OCR.
Currently, there are several limitations with this technique. At room temperature, the machine heats up to 31 &#176;C&#160;(this is the minimal measuring temperature while the machine is at room temperature), which may represent a stress state for the fly heads25. This however can be overcome by placing the machine in a cold room, which will enable measurements at 25 &#176;C&#160;and hence without a possible heat stress to the fly heads. Recent report has demonstrated placing the machine at 11 &#176;C, thus enabling the OCR recording of flies at 25 &#176;C21. Nevertheless, the fly head separation should be performed at room temperature. Furthermore, temperature fluctuations make it challenging to control pH changes and therefore it is highly recommended to test the impact of physiological conditions/drugs on OCR by using similar experimental setups. In addition, the contribution of oxygen consumption by non-mitochondrial-independent mechanisms has not yet been established26. By using various respiratory inhibitors that are efficient in fly heads, it would be possible to establish such non-mitochondrial oxygen consumption rates.
It is noteworthy that various mammalian maladies are characterized by alterations in energy metabolism. Among them are diseases that are characterized by either metabolic reduction such as Alzheimer&#39;s disease or metabolic rewiring such as cancer. Interestingly, KDAC inhibitors are used for both Alzheimer&#39;s disease and cancer treatment27. While the precise mechanisms by which KDAC inhibitors are achieving the therapeutic aspect remain unclear, the data from our technique supports the novel notion that such inhibitors may modulate metabolism.
In summary, this method is valuable for measuring overall oxygen consumption rates in vivo and more accurately displays drug effects on general metabolism, which may be overlooked in isolated mitochondria protocols12. For example, results obtained from this method, rather than previous techniques, have implicated novel insights for age-associated metabolic inflexibility upon KDAC treatment. While additional work is necessary to optimize the experimental conditions for fly heads, the combination of our technique and suitable analysis may lead to further elucidation of the mitochondrial activity in the context of whole living tissues.

Regulated metabolic activity is essential for the survival of cells and healthy function of a tissue. Deregulated metabolic activity has been extensively shown to be linked to the onset and progression of various maladies1. For example, lower metabolic activity was previously described in neurodegenerative diseases such as Alzheimer&#39;s and age-associated memory impairment2,3. Furthermore, mitochondrial dysfunction is believed to be causally involved in the aging process4,5. On the other hand, higher mitochondrial and metabolic rates were described in cancer cells6, where the use of mitochondrial inhibitors reduced tumorigenesis7.
One readout of metabolic activity is the oxygen consumption rate (OCR) of mitochondria. Interestingly, this type of readout is primarily obtained from isolated mitochondria or cells, thus the majority of what is described in the literature is mainly based on a readout that does not resemble the physiological state. However, there are several drawbacks to this technique. First, the protocol of mitochondrial isolation can potentially damage its integrity8, which may be a relevant artefact when comparing mitochondria isolated from young versus older tissues9. Furthermore, the isolation process is long and may result in loss of relevant protein posttranslational modifications which regulate mitochondrial function9,10,11. Moreover, it has been shown that isolated mitochondria do not consistently represent whole tissue metabolic rates12,13. Such cellular biological complexity could be viewed as, &#39;the whole is greater than the sum of its parts&#39;, i.e., mitochondria may display different metabolic rates inside a complex cell compared with their metabolic rate when isolated.
While cells may offer a better OCR readout than isolated mitochondria, cell to cell communication in the context of a whole tissue may be lost. For example, in the brain, the metabolic activity of neurons is highly dependent on the metabolic activity of neighboring glial cells14. As such, establishing new techniques to investigate OCR in whole tissue or whole organisms may prove more insightful for the onset and progression of various disorders.
Recently, new techniques have emerged to address these issues and enable the measurement of OCR from whole tissue, segment, or living organisms. For example, a recent work reported the oxygen measurement from a beetle flight muscle by using a permeabilized fiber approach with a respirometer15. New machines for micro-respirometry allow the measurement of OCR of pancreatic islets16,17. Consequently, it has been reported that this technology enables the measurement of OCR from whole worms18 and Zebra fish19. However, the presence of the digestive barrier may pose a challenge for testing various drugs in the context of OCR alterations. Interestingly, recent reports by Neville and colleagues have shown a new technique for measuring single drosophila larva brain with the well plate20,21.
In this study, we have used a similar setup to enable the measurement of whole OCR from whole living and non-mobile Drosophila22. This technique also offers a secondary advantage in measuring the impact of various drugs on metabolic activity in a whole segment, without having to pass through the digestive system barrier13,22. For example, it was previously demonstrated that direct injection of lysine deacetylase inhibitor (KDACi), a drug believed to alter epigenetic mechanism in the brain, resulted in an improved memories formation23. However, by using our novel technique, we discovered that KDAC inhibition resulted in a rapid increase of OCR, which may be a contributing factor by itself in the neuronal activity. Our protocol provides a simple and novel method to assess the impact of various drugs, genetic manipulation, or physiological states (disease, aging) on OCR in the context of a whole head.

<table><tbody><tr><td>Glucose</td><td>Sigma-Aldrich</td><td>G8644</td><td>D-(+)-Glucose solution 100 g/L in H2O, sterile-filtered</td></tr><tr><td>XF assay Medium</td><td>Agilent</td><td>103575-100</td><td>Seahorse XF DMEM Medium, pH 7.4</td></tr><tr><td>Sodium butyrate</td><td>Merck</td><td>817500</td><td>Dissolved in XF assay buffer</td></tr><tr><td>Seahorse XF24/e24 analyzer</td><td>Agilent</td><td></td><td></td></tr><tr><td>XF24/e24 Extracellular Assay Kit</td><td>Agilent</td><td>100850-001</td><td>Cartridge</td></tr><tr><td>XF24/e24 Islet Capture Microplates</td><td>Agilent</td><td>101122-100</td><td>Plate</td></tr><tr><td>Seahorse Capture Screen Insert Tool</td><td>Agilent</td><td>101135-10</td><td>Insertor</td></tr><tr><td>Petri dish</td><td>Sarstedt</td><td>821,472</td><td>Petri dish 92 x 16 mm</td></tr></tbody></table>

measuring and interpreting oxygen consumption rates in whole fly head segments

Regulated metabolic activity is essential for the normal functioning of living cells. Indeed, altered metabolic activity is causally linked with the progression of cancer, diabetes, neurodegeneration, and aging to name a few. For instance, changes in mitochondrial activity, the cell&#39;s metabolic powerhouse, have been characterized in many such diseases. Generally, the oxygen consumption rates of mitochondria were considered a reliable readout of mitochondrial activity and measurements in some of these studies were based on isolated mitochondria or cells. However, such conditions may not represent the complexity of a whole tissue. Recently, we have developed a novel method that enables the dynamic measurement of oxygen consumption rates from whole isolated fly heads. By utilizing this method, we have recorded lower oxygen consumption rates of the whole head segment in young versus aged flies. Secondly, we have discovered that lysine deacetylase inhibitors rapidly alter the oxygen consumption in the whole head. Our novel technique may therefore aid in uncovering new properties of various drugs, which may impact metabolic rates. Furthermore, our method may give a better understanding of metabolic behavior in an experimental setup that more closely resembles physiological states.

The ability to record high quality OCR measurement relies on centering the head in the middle of the net (Figure 1). This is important for the XF24 machine, which has a rather small oxygen sensor spot compared to the newer XFe24 machine in which the sensor is larger. As previously shown, centering the heads display a steady OCR for at least 20 consecutive measurements in young flies13.
One critical aspect of using the machines is to apply the correct analysis. It is recommended to check the oxygen levels during the experiments. Each 2 min measurement is subdivided into 10 sub-measurements (ticks). A well with 16 healthy heads usually displays an oxygen partial pressure (pO2) of 140-170 (mmHg) for the first tick. In the first example, we compared young vs. midlife fly heads (Figure 2A and 2B). While the oxygen levels drop quicker in the middle-aged heads, the observed difference is small (Figure 2A). Moreover, the range of the oxygen levels is similar between the conditions, with 165 during the first tick to 120 during the last tick. In such a case, it is preferable to use the AKOS algorithm to automatically generate the OCR (pmol/min)2, which reliably mirrors the oxygen level drop between young versus midlife heads (Figure 2B). Of note, the analysis program by the machine automatically chooses the AKOS algorithm.
However, based on our observations, automatically using the AKOS algorithm may give misleading, if not opposite results for the correct OCR. Such artefacts can be generated in conditions of a highly consuming sample which reaches anoxia13,22. For example, the addition of sodium butyrate (SB), a KDAC inhibitor, transiently changes the dynamics of the oxygen levels (Figure 3A). Whereas the vehicle controls display steady levels of oxygen during the first and last ticks, SB addition causes a considerable and transient drop of the oxygen levels in these ticks (Figure 3A). SB by itself does not alter the oxygen levels in the background wells, where no heads are added (data not shown). The data supports the notion that SB increases oxygen consumption. As the collection of first tick is delayed (12 seconds until the first tick is recorded in the measuring phase) the first data point is already lower in the SB treated wells. Therefore, it is difficult to capture the early changes in oxygen consumption following the addition of this HDAC inhibitor. Furthermore, the oxygen levels in the SB treated samples are reduced to already low levels (anoxia) as indicated by the collection of the last ticks. At anoxia, the heads slow down their oxygen consumption in the last ticks (Figure 3A). Because the AKOS calculation takes into account all ticks and ignores an anoxic state, it generates a misleading OCR. Indeed, the non-normalized AKOS based OCR levels show little change upon the injection (dashed line) of port A (Veh/SB) (Figure 3B).
Normalizing the OCR levels to the pre-injection measurement based on the AKOS reveals very similar levels of OCR before and after the injection of port A, which does not support the oxygen level changes (Figure 3A). Under these circumstances, the FIXED algorithm, which more closely models/resembles the OCR and oxygen level changes is recommended (Figure 3C). Consequently, the FIXED algorithm based normalized measurement reveals an increased OCR upon SB treatment (Figure 3C).
A drawback with the new machine is the absence of the FIXED algorithm. Therefore, in experiments where highly consuming sample/treatment is used, it is recommended to calculate the OCR measurements manually, and calculating the decrease in oxygen level per time for the first 3-5 ticks in each measurement.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/58601/58601fig1.jpg" /> 
Figure 1. An example of a well containing 16 one-week old heads of male flies. The heads are centered below a net and floating in the media. <a href="https://www.jove.com/files/ftp_upload/58601/58601fig1large.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/58601/58601fig2.jpg" /> 
Figure 2. A representative example of OCR measurement comparison between one-week old fly heads (young) and four-week old fly heads (middle-age). (A) The oxygen levels are shown for three separate measurements; each 2 min measurement is subdivided into ten sub-measurements (ticks). (B) A quantification of (A). The levels of the first and last ticks are similar, although the levels of the middle-age sample are slightly lower. A quantification of the slope of the decrease in oxygen levels is used to generate the OCR levels. As previously described22, the OCR of middle aged flies is 10%-15% higher than young flies. <a href="https://www.jove.com/files/ftp_upload/58601/58601fig2large.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/58601/58601fig3.jpg" /> 
Figure 3. An example of the alteration of OCR by sodium butyrate (SB) in young fly heads. (A) Oxygen levels recorded from seven measurements following the addition of 15 mM SB. The dashed line marks the injection of the drug (or vehicle) from port A. Of note, while the oxygen levels of ticks 1 and 10 remain stable in the control group (Blue), the levels of oxygen during these ticks are transiently (six measurements following the injection) reduced in the SB treated samples (orange). In addition, the decrease in oxygen levels is greatly reduced during the last ticks of SB treated samples. N=3 per group (B) [Left] Non-normalized OCR levels calculated from the AKOS algorithm. The calculation incorrectly shows similar levels of OCR before and after the injection of SB by port A. [Right] Normalization of the OCR to the measurement prior to the injection of port A. (C) [Left] &#39;Fixed&#39; algorithm calculation of (A) showing the non-normalized OCR. Here, the OCR closely represents the transient increase in oxygen usage of the heads upon SB treatment; [Right] Normalization of the OCR to the measurement prior to the injection of port A. Error bars indicate the S.E.M. in all the graphs. <a href="https://www.jove.com/files/ftp_upload/58601/58601fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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Measuring and Interpreting Oxygen Consumption Rates in Whole Fly Head Segments

Measuring alterations in metabolic rates is central to understanding the progression of various diseases and aging. Here, we present a novel technique to measure whole head oxygen consumption that more closely resembles the physiological state and may aid in revealing novel drugs that modify mitochondrial activity.

Regulated metabolic activity is essential for the normal functioning of living cells. Indeed, altered metabolic activity is causally linked with the ...

measuring-interpreting-oxygen-consumption-rates-whole-fly-head

Research

JoVE Journal

Biochemistry

15.6K Views.  Ludwig-Maximilians University of Munich. Deze methode kan helpen bij het beantwoorden van belangrijke vragen op het gebied van metabolisme, zoals, hoe metabole activiteit wordt veranderd tijdens het proces van veroudering. Het belangrijkste voordeel van deze techniek is het gebruik van een intact lichaamssegment dat gegevens heeft die sterk lijken op de natuurlijke en fysiologische toestand in vergelijking met de klassieke mitochondriale isolatie. Het aantonen van de procedure zal Anuroop en Annika, een grad student in mijn laborato...

Video: Measuring and Interpreting Oxygen Consumption Rates in Whole Fly Head Segments

Identification of Small Molecule-binding Proteins in a Native Cellular Environment by Live-cell Photoaffinity Labeling

Identifying the molecular target(s) of small molecules is a challenging but necessary step towards understanding their mechanism of action. While several target identification methods have been developed and used to successfully elucidate the binding proteins of a variety of small molecules, these techniques have drawbacks that make them unsuitable for detecting certain types of small molecule-target interactions. In particular, non-covalent interactions that depend on native cellular conditions, such as those of membrane proteins whose structures may be perturbed upon cell lysis, are often not amenable to affinity-based target identification methods. Here, we demonstrate a method wherein a probe containing a photolabile group is used to covalently crosslink to the small molecule binding protein within the environment of the live cell, allowing the detection and isolation of the target protein without the need for maintenance of the interaction after cell lysis. This technique is a valuable tool for studying biologically interesting small molecules with unknown mechanisms, both in the context of basic biology as well as drug discovery.

We describe here a method for identification of small molecule-binding proteins using photoaffinity labeling. The advantage of this technique is that binding and covalent labeling of the target proteins occurs within the live cellular environment, removing the risk of disrupting native protein structure and binding conditions upon cell lysis.

Identifying the molecular target(s) of small molecules is a challenging but necessary step towards understanding their mechanism of action. While several ...

A Method for Measuring RNA N6-methyladenosine Modifications in Cells and Tissues

N6-Methyladenosine (m6A) modifications of RNA are diverse and ubiquitous amongst eukaryotes. They occur in mRNA, rRNA, tRNA, and microRNA. Recent studies have revealed that these reversible RNA modifications affect RNA splicing, translation, degradation, and localization. Multiple physiological processes, like circadian rhythms, stem cell pluripotency, fibrosis, triglyceride metabolism, and obesity are also controlled by m6A modifications. Immunoprecipitation/sequencing, mass spectrometry, and modified northern blotting are some of the methods commonly employed to measure m6A modifications. Herein, we present a northeastern blotting technique for measuring m6A modifications. The current protocol provides good size separation of RNA, better accommodation and standardization for various experimental designs, and clear delineation of m6A modifications in various sources of RNA. While m6A modifications are known to have a crucial impact on human physiology relating to circadian rhythms and obesity, their roles in other (patho)physiological states are unclear. Therefore, investigations on m6A modifications have immense possibility to provide key insights into molecular physiology.

A Method for Measuring RNA N6-methyladenosine Modifications in Cells and Tissues

A modified northern blotting method for measuring N6-methyladenosine (m6A) modifications in RNA is described. The current method can detect modifications in diverse RNAs and controls under various experimental designs.

N6-Methyladenosine (m6A) modifications of RNA are diverse and ubiquitous amongst eukaryotes. They occur in mRNA, rRNA, tRNA, and microRNA. Recent studies ...

Measurement of Mitochondrial Oxygen Consumption in Permeabilized Fibers of Drosophila Using Minimal Amounts of Tissue

The fruit fly, Drosophila melanogaster, represents an emerging model for the study of metabolism. Indeed, drosophila have structures homologous to human organs, possess highly conserved metabolic pathways and have a relatively short lifespan that allows the study of different fundamental mechanisms in a short period of time. It is, however, surprising that one of the mechanisms essential for cellular metabolism, the mitochondrial respiration, has not been thoroughly investigated in this model. It is likely because the measure of the mitochondrial respiration in Drosophila usually requires a very large number of individuals and the results obtained are not highly reproducible. Here, a method allowing the precise measurement of mitochondrial oxygen consumption using minimal amounts of tissue from Drosophila is described. In this method, the thoraxes are dissected and permeabilized both mechanically with sharp forceps and chemically with saponin, allowing different compounds to cross the cell membrane and modulate the mitochondrial respiration. After permeabilization, a protocol is performed to evaluate the capacity of the different complexes of the electron transport system (ETS) to oxidize different substrates, as well as their response to an uncoupler and to several inhibitors. This method presents many advantages compared to methods using mitochondrial isolations, as it is more physiologically relevant because the mitochondria are still interacting with the other cellular components and the mitochondrial morphology is conserved. Moreover, sample preparations are faster, and the results obtained are highly reproducible. By combining the advantages of Drosophila as a model for the study of metabolism with the evaluation of mitochondrial respiration, important new insights can be unveiled, especially when the flies are experiencing different environmental or pathophysiological conditions.

In this paper, a method to measure oxygen consumption using high-resolution respirometry in permeabilized thoraxes of Drosophila is described. This technique requires a minimal amount of tissue compared to the classic mitochondrial isolation technique and the results obtained are more physiologically relevant.

The fruit fly, Drosophila melanogaster, represents an emerging model for the study of metabolism. Indeed, drosophila have structures homologous to human ...

A Colorimetric Method for Measuring Iron Content in Plants

Iron, one of the most important micronutrients in living organisms, is involved in basic processes, such as respiration and photosynthesis. Iron content is rather low in all organisms, amounting in plants to about 0.009% of dry weight. To date, one of the most accurate methods for measuring iron concentration in plant tissues is flame absorption atomic spectroscopy. However, this approach is time-consuming and expensive and requires specific equipment not commonly found in plant laboratories. Therefore, a simpler, yet accurate method that can be routinely used is needed. The colorimetric Prussian Blue method is regularly used for qualitative iron staining in animal and plant histological sections. In this study, we adapted the Prussian Blue method for quantitative measurements of iron in tobacco leaves. We validated the accuracy of this method using both atomic spectroscopy and Prussian Blue staining to measure iron content in the same samples and found a linear regression (R2 = 0.988) between the two procedures. We conclude that the Prussian Blue method for quantitative iron measurement in plant tissues is precise, simple, and inexpensive. However, the linear regression presented here may not be appropriate for other plant species, due to potential interactions between the sample and the reagent. Establishment of a regression curve is thus needed for different plant species.

We present a simple and reliable protocol for measuring iron content in plant tissues using the colorimetric Prussian Blue method.

Iron, one of the most important micronutrients in living organisms, is involved in basic processes, such as respiration and photosynthesis. Iron content ...

Sampling and Pretreatment of Tooth Enamel Carbonate for Stable Carbon and Oxygen Isotope Analysis

Stable carbon and oxygen isotope analysis of human and animal tooth enamel carbonate has been applied in paleodietary, paleoecological, and paleoenvironmental research from recent historical periods back to over 10 million years ago. Bulk approaches provide a representative sample for the period of enamel mineralization, while sequential samples within a tooth can track dietary and environmental changes during this period. While these methodologies have been widely applied and described in archaeology, ecology, and paleontology, there have been no explicit guidelines to aid in the selection of necessary lab equipment and to thoroughly describe detailed laboratory sampling and protocols. In this article, we document textually and visually, the entire process from sampling through pretreatment and diagenetic screening to make the methodology more widely available to researchers considering its application in a variety of laboratory settings.

Stable carbon and oxygen isotope analysis of human and animal tooth enamel carbonate has been used as a proxy for individual diet and environmental reconstruction. Here, we provide a detailed description and visual documentation of bulk and sequential tooth enamel sampling as well as pretreatment of archaeological and paleontological samples.

Stable carbon and oxygen isotope analysis of human and animal tooth enamel carbonate has been applied in paleodietary, paleoecological, and ...

An Affordable and Efficient "Homemade" Platform for Drosophila Behavioral Studies, and an Accompanying Protocol for Larval Mitochondrial Respirometry

The usefulness of Drosophila as a model organism for the study of human diseases, behaviors and basic biology is unquestionable. Although practical, Drosophila research lacks popularity in developing countries, possibly due to the misinformed idea that establishing a lab and performing relevant experiments with such tiny insects is difficult and requires expensive, specialized apparatuses. Here, we describe how to build an affordable flylab to quantitatively analyze a myriad of behavioral parameters in D. melanogaster, by 3D-printing many of the necessary pieces of equipment. We provide protocols to build in-house vial racks, courtship arenas, apparatuses for locomotor assays, etc., to be used for general fly maintenance and to perform behavioral experiments using adult flies and larvae. We also provide protocols on how to use more sophisticated systems, such as a high resolution oxygraph, to measure mitochondrial oxygen consumption in larval samples, and show its association with behavioral changes in the larvae upon the xenotopic expression of the mitochondrial alternative oxidase (AOX). AOX increases larval activity and mitochondrial leak respiration, and accelerates development at low temperatures, which is consistent with a thermogenic role for the enzyme. We hope these protocols will inspire researchers, especially from developing countries, to use Drosophila to easily combine behavior and mitochondrial metabolism data, which may lead to information on genes and/or environmental conditions that may also regulate human physiology and disease states.

An Affordable and Efficient "Homemade" Platform for Drosophila Behavioral Studies, and an Accompanying Protocol for Larval Mitochondrial Respirometry

We provide protocols for anyone with a "maker culture" mind to start building a flylab for quantitative analysis of a myriad of behavioral parameters in Drosophila melanogaster, by 3D-printing many of the necessary pieces of equipment. We also describe a high resolution respirometry protocol using larvae to combine behavioral and mitochondrial metabolism data.

The usefulness of Drosophila as a model organism for the study of human diseases, behaviors and basic biology is unquestionable. Although practical, ...

Behavior

Metabolic Analysis of Drosophila melanogaster Larval and Adult Brains

This protocol describes a method for measuring the metabolism in Drosophila melanogaster larval and adult brains. Quantifying metabolism in whole organs provides a tissue-level understanding of energy utilization that cannot be captured when analyzing primary cells and cell lines. While this analysis is ex vivo, it allows for the measurement from a number of specialized cells working together to perform a function in one tissue and more closely models the in vivo organ. Metabolic reprogramming has been observed in many neurological diseases, including neoplasia, and neurodegenerative diseases. This protocol was developed to assist the D. melanogaster community&#39;s investigation of metabolism in neurological disease models using a commercially available metabolic analyzer. Measuring metabolism of whole brains in the metabolic analyzer is challenging due to the geometry of the brain. This analyzer requires samples to remain at the bottom of a 96-well plate. Cell samples and tissue punches can adhere to the surface of the cell plate or utilize spheroid plates, respectively. However, the spherical, three-dimensional shape of D. melanogaster brains prevents the tissue from adhering to the plate. This protocol requires a specially designed and manufactured micro-tissue restraint that circumvents this problem by preventing any movement of the brain while still allowing metabolic measurements from the analyzer&#39;s two solid-state sensor probes. Oxygen consumption and extracellular acidification rates are reproducible and sensitive to a treatment with metabolic inhibitors. With a minor optimization, this protocol can be adapted for use with any whole tissue and/or model system, provided that the sample size does not exceed the chamber generated by the restraint. While basal metabolic measurements and an analysis after a treatment with mitochondrial inhibitors are described within this protocol, countless experimental conditions, such as energy source preference and rearing environment, could be interrogated.

Metabolic Analysis of Drosophila melanogaster Larval and Adult Brains

We present a protocol for measuring oxygen consumption and extracellular acidification in Drosophila melanogaster larval and adult brains. A metabolic analyzer is utilized with an adapted and optimized protocol. Micro-tissue restraints are a critical component of this protocol and were designed and created specifically for their use in this analysis.

This protocol describes a method for measuring the metabolism in Drosophila melanogaster larval and adult brains. Quantifying metabolism in whole organs ...

Neuroscience

Measurement of Metabolic Rate in Drosophila using Respirometry

Metabolic disorders are a frequent problem affecting human health. Therefore, understanding the mechanisms that regulate metabolism is a crucial scientific task. Many disease causing genes in humans have a fly homologue, making Drosophila a good model to study signaling pathways involved in the development of different disorders. Additionally, the tractability of Drosophila simplifies genetic screens to aid in identifying novel therapeutic targets that may regulate metabolism. In order to perform such a screen a simple and fast method to identify changes in the metabolic state of flies is necessary. In general, carbon dioxide production is a good indicator of substrate oxidation and energy expenditure providing information about metabolic state. In this protocol we introduce a simple method to measure CO2 output from flies. This technique can potentially aid in the identification of genetic perturbations affecting metabolic rate.

Measurement of Metabolic Rate in Drosophila using Respirometry

Metabolic disorders are among one of the most common diseases in humans. The genetically tractable model organism D. melanogaster can be used to identify novel genes that regulate metabolism. This paper describes a relatively simple method which allows studying the metabolic rate in flies by measuring their CO2 production.

Metabolic disorders are a frequent problem affecting human health. Therefore, understanding the mechanisms that regulate metabolism is a crucial ...

Biology

Adapting Microrespirometry for Drosophila - Insights into CASK Mutant Metabolism

Measuring O2 Consumption in Drosophila melanogaster Using Coulometric Microrespirometry

Coulometric microrespirometry is a straightforward, inexpensive method for measuring the O2 consumption of small organisms while maintaining a stable environment. A coulometric microrespirometer consists of an airtight chamber in which O2 is consumed and the CO2 produced by the organism is removed by an absorbent medium. The resulting pressure decrease triggers electrolytic O2 production, and the amount of O2 produced is measured by recording the amount of charge used to generate it. In the present study, the method has been adapted to Drosophila melanogaster tested in small groups, with the sensitivity of the apparatus and the environmental conditions optimized for high stability. The amount of O2 consumed by wildtype flies in this apparatus is consistent with that measured by previous studies. Mass-specific O2 consumption by CASK mutants, which are smaller and known to be less active, was not different from congenic controls. However, the small size of CASK mutants resulted in a significant reduction in O2 consumption on a per-fly basis. Therefore, the microrespirometer is capable of measuring O2 consumption in D. melanogaster, can distinguish modest differences between genotypes, and adds a versatile tool for measuring metabolic rates.

Author Spotlight: Influence of Temperature on Drosophila melanogaster and Desert-Adapted Beetles 

Coulometric respirometry is ideal for measuring the metabolic rate of small organisms. When adapted for Drosophila melanogaster in the present study, measured O2 consumption was within the range reported for wildtype D. melanogaster by previous studies. Per-fly O2 consumption by CASK mutants, which are smaller and less active, was significantly lower than the wildtype.

Coulometric microrespirometry is a straightforward, inexpensive method for measuring the O2 consumption of small organisms while maintaining a stable ...

Measurement of Oxygen Consumption Rates in Intact Caenorhabditis elegans

Optimal mitochondrial function is critical for healthy cellular activity, particularly in cells that have high energy demands like those in the nervous system and muscle. Consistent with this, mitochondrial dysfunction has been associated with a myriad of neurodegenerative diseases and aging in general. Caenorhabditis elegans have been a powerful model system for elucidating the many intricacies of mitochondrial function. Mitochondrial respiration is a strong indicator of mitochondrial function and recently developed respirometers offer a state-of-the-art platform to measure respiration in cells. In this protocol, we provide a technique to analyze live, intact C. elegans. This protocol spans a period of ~7 days and includes steps for (1) growing and synchronization of C. elegans, (2) preparation&#160;of compounds to be injected and hydration of probes, (3) drug loading and cartridge equilibration, (4) preparation of worm assay plate and assay run, and (5) post-experiment data analysis.

Measurement of Oxygen Consumption Rates in Intact Caenorhabditis elegans

Mitochondrial respiration is critical for organismal survival; therefore, oxygen consumption rate is an excellent indicator of mitochondrial health. In this protocol, we describe the use of a commercially available respirometer to measure basal and maximal oxygen consumption rates in live, intact, and freely-motile Caenorhabditis elegans.

Optimal mitochondrial function is critical for healthy cellular activity, particularly in cells that have high energy demands like those in the nervous ...

Measuring and Interpreting Oxygen Consumption Rates in Whole Fly Head Segments

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