M.Z. and K.S.F. and this work was supported by a grant awarded under the Pitt Innovation Challenge (PInCh), through the Clinical and Translational Science Institute at the University of Pittsburgh, and NHLBI R01 grant HL153407, awarded to M.Z.

The University of Pittsburgh&#8217;s Institutional Animal Care and Use Committee approved all animal protocols specified in this publication prior to undertaking any of these animal experiments.
NOTE: This protocol details how to perform in vivo mucociliary clearance studies utilizing radionuclide imaging with a dual-modality SPECT/CT scanner. The techniques demonstrated are running system calibrations, anesthetizing mice, tracheal intubation of mice, instilling the isotope into the lungs, dual-modality imaging, co-registration of these images, and analysis.
1. SPECT/CT system setup
<ol>
	<li>Design an appropriate workflow and set up prior to running experiments using living animals.
	<ol>
		<li>Use a SPECT acquisition consisting of 60 projections with a step size of 6o between projections with a 40 cm radius of rotation. The CT acquisition consists of 220 projections with a 1.6o angle between projections.</li>
	</ol>
	</li>
	<li>Ensure that the system has the correct MWB collimators for mice and SPECT imaging in place. If the inappropriate collimators are installed, use the collimator wizard to install the correct ones.</li>
	<li>Run the necessary system calibrations to prepare the system for use. 
	NOTE: The SPECT and CT components of the scanner need calibration. Calibrate the CT components using a source conditioning and a Dark/Light (D/L) calibration once a day, a Center Offset (COS) calibration every 2 weeks, and evaluate the x-ray hardware every month. The SPECT components need to be calibrated once a year.
	<ol>
		<li>To evaluate the x-ray hardware, check the Evaluate X-ray hardware box during CT calibrations (Supplemental CT Calibration Menu).</li>
		<li>To perform source conditioning, check the Perform source conditioning box during CT calibrations (Supplemental CT Calibration Menu).</li>
		<li>To perform a D/L calibration, check the D/L box next to the CT acquisition protocol used during experiments during CT calibrations. Uncheck all other protocols (Supplemental CT Calibration Menu).</li>
		<li>To perform a COS calibration, replace the bed with the calibration ring tool, adjust the bed type settings to match in the motion control settings, and check the COS box next to the CT acquisition protocol used during experiments during CT calibrations. Uncheck all other protocols (Supplemental CT Calibration Menu, Supplemental Calibration Ring).</li>
	</ol>
	</li>
</ol>
2. Mouse Intubation and Instillation
<ol>
	<li>Weigh the mice to be scanned. If scanning multiple mice, take care to mark the mice for identification purposes using methods such as ear punching or marking of the tail.</li>
	<li>Anesthetize a mouse using 1.5% isoflurane with a gas flow of 2 L/min O2 in a gas chamber for ~5 minutes to produce anesthesia of sufficient depth, until breathing slows to ~55-65 breaths per minute 16 (Figure 1A).</li>
	<li>Remove the mouse from the chamber and suspend by the front incisors on an intubation stand at a 45o incline. Equip the intubation stand with a nose cone to ensure the mouse is anesthetized during the intubation (Figure 1B).</li>
	<li>Connect one end of a 50 &#181;m fiber optic wire to a light source and thread a 20-gauge cannula over it using the wire to act as a guide (Figure 1C).</li>
	<li>Open the mouth of the mouse and pull the tongue forward using blunt forceps. Illuminate the guide wire and use it to visualize the vocal cords (Figure 1D).</li>
	<li>Pass the guide wire through the vocal cords so that the wire is just beyond the vocal cords and resting in the upper trachea. Slide the 1 inch cannula forward along the wire to intubate the mouse, passing the cannula deep enough so that the hub of it is against the animal&#8217;s incisors (Figure 1E). Remove the wire leaving the cannula in place.</li>
	<li>Test the intubation by briefly plugging the cannula with a finger and checking for changes in breathing. Halted breathing or strained breathing while plugging and accelerated breathing upon release are signs of proper tracheal intubation. If there is no change in respiratory patterns upon plugging the cannula, the latter is likely in the esophagus.</li>
	<li>Prepare 0.2 mCi of 99mtechnetium-sulfur colloid (99mTc-Sc) in a volume of 10 &#181;L, and pipette into the cannula. Allow the mouse to spontaneously inhale it into the lungs over 1-2 min (Figure 1F). Remove the cannula before transferring the mouse to the pallet of the scanner. 
	NOTE: The radionuclide was prepared and filtered by Cardinal Health.</li>
</ol>
3. SPECT/CT Imaging
<ol>
	<li>Transfer the mouse to a 25 mm pallet with a nose cone and secure with tape, taking care to not tape the chest and abdomen too tightly to avoid impairing respiration. Take care to remove any metal ear tags attached to the mouse.</li>
	<li>Prepare a radioactive phantom consisting of 0.05 mCi in 200 &#181;L and place this amount in a 0.2 mL PCR tube. Position the tube by taping to the pallet under the lower abdomen of the mouse, avoiding overlap with the lungs. 
	NOTE: The phantom is used for the purpose of co-registering CT and SPECT images, as well as a negative control for clearance.</li>
	<li>Insert the mouse into the SPECT/CT system, select the imaging workflow, and run Setup.</li>
	<li>Set up the positioning of the detectors on the mouse, and run the imaging workflow.</li>
	<li>Prepare a cage for mice that have received radioactivity post-procedure, with unrestricted access to food and water, and clear labeling using a radiation safety sticker.</li>
	<li>Upon completion of the workflow, remove the mouse from the imaging pallet, and allow it to recover in the prepared cage for a duration of 6 hours between scans (end of scan 1 to beginning of scan 2) with ad libitum access to food and water. 6 h was chosen as it corresponds to the time period in which linear clearance depending on cilia function is taking place with very little alveolar clearance.</li>
	<li>After 6 hours, re-anaesthetize the mouse and scan, along with the phantom, using the same workflow to measure the amount of isotope cleared from the airways. 
	NOTE: It is critical to allow the mouse to recover as uninterrupted anesthesia with isoflurane for 6 hours will lead to a significant cilia-depressant effect, resulting in near zero mucociliary clearances.</li>
</ol>
4. Analysis
<ol>
	<li>After imaging, perform post-processing to reconstruct complete 3D stack images.
	<ol>
		<li>Histogram the SPECT images using the factory standard settings for 99mTc, and then reconstruct using a MAP3D algorithm and point spread function (PSF) reconstruction. 
		NOTE: The reconstruction was performed using 8 iterations and 6 subsets. An effective reconstruction needs a ratio of subsets to projections at 1:10 or divide evenly into the number of projections, so 6 subsets were used due to the acquisition using 60 projections.</li>
		<li>Reconstruct the CT images using the Feldkamp algorithm and a Shepp-Logan filter. 
		NOTE: The reconstruction was performed using 4 iterations.</li>
	</ol>
	</li>
	<li>Process the CT and SPECT images in FIJI ImageJ21 using the reslice tool to generate coronal view images from the default axial images. Then perform a z-stack sum projection on the SPECT image to add the count data from each slice and generate a single image for ease of analysis.</li>
	<li>Resize and co-register the CT and SPECT images using the phantom Eppendorf tube as a reference (Figure 2A,B). Track and use consistent resize measurements across all samples.</li>
	<li>Binarize the CT image using auto thresholding, followed by inverting the stack, and performing a z-stack sum projection to generate an outline of the lungs for analysis (Figure 2C).</li>
	<li>Rotate the CT and SPECT images and merge the image using the channel tools. Calculate MCC by drawing an ROI around the right lung and measuring (Figure 2D). 
	NOTE: This measurement will be of the total counts in the right lung for the 0 and 6 hour time points, with the 6 hour images corrected for radioactive decay using the formula: N(t) = N0e&#8722;t. 99mTc-Sc has a decay constant of 3.21e&#8722;5 per second with a half-life of ~6 hours. These values can then be used to calculate a percent clearance. 
	NOTE: Right lung is chosen for ROI drawing and measuring counts as the mucociliary clearance will transport the radioisotope out of the lungs to the pharynx from where it will be swallowed and end up in the stomach. Quite frequently, counts can be seen in the stomach that may overlap with the left lung and hence produce erroneous counts. This confounding can be avoided by measuring counts in the right lung only.</li>
</ol>

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None related to this work.

The role of motile respiratory cilia in both disease and development continues to evolve and be better appreciated. Synchronous, metachronal beating of multiple motile cilia on the apical surface of cells lining the tracheobronchial tree generate cephalad flow producing mucociliary clearance or MCC. MCC is compromised in ciliopathies like PCD22, acquired diseases like COPD18, and its importance is being recognized in CHDs, not traditionally considered to be ciliopathies. Recent data has shown respiratory ciliary dysfunction in both CHD with heterotaxy23 and without heterotaxy7. Such motile cilia dysfunction was shown to translate into greater respiratory symptoms9 as well as greater post-operative morbidity8. Most, if not all, of these diseases, have mouse models available and our protocol for measuring MCC in mice is a valuable tool that can be utilized to test potential therapeutics.
Animal models provide utility for understanding diseases and the development of therapies. In vivo animal imaging provides further utility with the ability to acquire multiple data points from the same animals, without the need to sacrifice the animals, allowing investigators to follow longitudinal course of disease as well as study duration of treatment effects. The mouse model of MCC has been developed over the course of decades by multiple investigators, initially being performed on beagle dogs using planar scintigraphy, a two dimensional nuclear imaging technique24. The technique was adapted for use in mice a decade later, followed by adaptation to SPECT imaging a decade after that25,26. The development of this technique in mouse models was a major development in the relevance of this technique, due to the availability of multiple mouse models of human diseases like PCD in which ciliary function is significantly altered. MCC has been assessed in mouse models of lung denervation and immunosuppression, and has the potential to be used in conjunction with other models19,26. MCC measurement studies in human patients with airway diseases such as CF, asthma, PCD, and ciliopathies associated with CHD have been conducted, and have yielded results that the technique can aid both studies of lung physiology and therapeutic efficacy13.
An important part of this protocol is setting up acquisitions with the correct imaging parameters to acquire accurate images for quantification. A number of factors are key when designing SPECT acquisition settings, including which collimators are used, the number of projections to acquire per revolution, and rotation step size. Collimator selection is a major factor in the sensitivity and resolution of the acquisition, and acquisition settings may need to be tailored to the collimator being used27. Alternately, when using bigger animals like rats, the collimators would need to be adjusted. Multiple pinhole collimators for example are more sensitive, but care should be taken when selecting a step size in order to avoid overlapping projections and causing undesired multiplexing, which can further increase sensitivity of the acquisition at the expense of some image ambiguity that can cause reconstruction artifacts25. Reconstruction setup is also key to generate quantifiable images. MAP3D is a commonly used iterative reconstruction algorithm, and PSF is a common reconstruction model. Both are reliable for reconstructing images, but care should be taken when setting the number of iterations and subsets. A higher number of iterations will increase the computational time required for the reconstruction, and increase the quality of the reconstruction with diminishing returns upon further increase.
In order to quantify images in ImageJ, the ideal measurement tool to use is RawIntDen, which outputs the sum value of pixels in a selection. When quantifying SPECT data across differently sized lung ROIs, the use of RawIntDen provides an absolute measure of counts and avoids adjusting the measurement to the area of the ROI, like the mean measurement would21.
This technique has a number of associated sources of error that the investigator should be cognizant of when applying this technique. A notable confounder is the use of anesthetic agents. Isoflurane is a fast acting, inhaled anesthetic that the mice recover from rapidly after completion of an acquisition. However, care should be taken to provide the mice with ample time to recover in their cages, and not kept anesthetized any longer than necessary. In our personal experience (unpublished data) mice that were kept anesthetized continuously using inhaled isoflurane between the 0 and 6 hour time-point showed negligible clearance. Likewise, a controlled dose of anesthetic is also necessary to ensure rapid recovery. When securing the animal to the pallet for imaging, the phantom tube used for co-registration should be kept low on the stomach to avoid artifacts from overlapping with the lungs. Likewise, to ensure a quality CT image, take care to remove any metal tags from the mouse to avoid artifacts from x-ray scattering.
The current MCC protocol can be applied to myriad animal models. This technique has a negligible effect on the health of the animal scanned, is well tolerated by mice, and because of this it can be used with disease models without risking the health of already delicate mice. The strength of this methodology comes from it being an in vivo technique, which allows for the acquisition of consistent and repeatable measurements of airway function without the sacrifice of animals to excise tracheas for video-microscopy, that ex vivo models require26. The consistency of this technique in producing repeatable measurements across multiple scans of the same animals, allows for the same animal to be treated with different agents or potential therapeutics, and statistical comparisons made between the same animal to reduce biological variability inherent in any animal model, thereby reducing the sample size needed to show statistically significant differences.
The assessment of airway function using the MCC technique can be adjusted to a variety of animal models and applied to many different models of airway health, as well as testing new therapies. The airways of mouse models of PCD can be assessed using this technique, as well as models of COPD. Our method can also be utilized to study differential effects of various anesthetics on MCC that are in common clinical use. Finally, the effects of therapeutic agents on the airway can also be assessed using this model. As previously stated but bears repetition, as it is an in vivo measurement it allows for repeat MCC assessments over the course of a disease, as well as test benefits of therapeutic interventions over time. Additionally, mice are the most common laboratory animals used to mimic/study human diseases, with, in some cases, multiple transgenic mouse models of human disease available to choose from.

Cilia are microtubule based cellular organelles conserved across evolutionary history from algae to humans. They emanate from cell surfaces and have a number of functions1, ranging from recognition of local environmental sensory signals to motility, functions which can be traced back from humans to early unicellular eukaryotic organisms2,3. Cilia can be non-motile and single serving as a cell&#8217;s specialized antenna to process environmental signals; or motile and multiple, beating in synchronized, metachronal waves to generate fluid flow, such as in the lining of the fallopian tubes and the upper and lower airways, except for the terminal bronchioles leading to the alveoli1,2.
The extensive epithelial surface of the respiratory tract is exposed to a constant barrage of contamination in the form of a variety of potentially hazardous inhaled pollutants and pathogens, necessitating a defense. One key defense mechanism is the mucociliary apparatus of the tracheobronchial tree, where a continuous flow of secreted mucus is mechanically transported out of the airway by the beating of multiple motile cilia lining the apical surfaces of the tracheo-bronchial epithelial cells. These function to entrap inhaled contaminants, and through their continuous, synchronal beating, transport them cephalad4,5.
Cilia have been demonstrated to have key roles such as in the development of left-right patterning in developing embryos, where motile cilia at the embryonic node break symmetry6. Mutations in cilia related genes have been linked to diseases such as congenital heart disease (CHD) due to the asymmetric structure of the heart6. Recent studies have reported a high incidence of ciliary dysfunction in the respiratory tracts of patients with CHD, as well as an increased prevalence of post-operative respiratory complications and chronic respiratory tract symptoms in the upper and lower airways7,8,9,10. Patients with CHD and ciliary dysfunction, with or without heterotaxy, have been demonstrated to have increased risk of respiratory complications and negative respiratory outcomes post-operatively5,8,10. Beyond their roles in signaling and development, the importance of airway cilia has been demonstrated by ciliopathies, of which a prime example is primary ciliary dyskinesia (PCD). PCD is a congenital disorder resulting from a number of mutations affecting the motile respiratory cilia, leading to recurrent lung infections, bronchiectasis, and potentially the need for lung transplantation11. Additionally, even though cilia are normal in cystic fibrosis (CF), most common congenital disorder in the Caucasian population, MCC is impaired due to thick, viscous mucus resulting from mutations in the CFTR gene12. There are multiple mouse models of PCD and CF, as well as an ever-increasing number of models of CHD. Ultimately cilia are versatile structures with many key roles, and a method to assess the function of motile respiratory cilia in vivo can be valuable for pre-clinical study, and assessing effects of mutations as well as drugs on mucociliary clearance (MCC)13. The method would also be valuable in assessing effects of novel drugs, gene therapy or interventions on MCC in these mouse models.
There are many different models that have been used to assess MCC. One notable method involves the use of methylene blue dye that has been instilled into the bronchus, with clearance measured by fiberoptic measurement of dye movement14. This method is limited by the ability to observe the movement of the dye, which is more routine in humans than in pre-clinical mouse models. Another notable method is synchrotron phase-contrast X-ray imaging (PCXI), which can be used to track individual particles in an airway. This method is relatively new and not widely accessible15. There are numerous ex vivo methods of assessing the airway by excising a trachea for video-microscopy, however these models provide little utility in human patients16. High resolution techniques for cilia imaging such as optical coherence tomography are limited in the same way17.
In this article, we present a reproducible method to measure MCC in vivo that has been used to measure lung clearances in myriad animal models, as well as study MCC in chronic obstructive pulmonary disease and assess the effects of immunosuppressive drugs18,19. This method tracks the clearance of the radiopharmaceutical 99mtechnetium-sulfur colloid (99mTc-Sc), an insoluble particulate radiotracer, after instillation into the lungs. The radionuclide can then be tracked using single photon emission computed tomography (SPECT)18,20. We have further refined this technique for measuring MCC by using dual modality SPECT and computed tomography (CT) imaging with co-localization of radioisotope counts to the lungs and measuring the decrease in these counts over 6 hours. Dual-modality imaging, with co-registration of CT and SPECT images allows for accurate localization of radiation counts to our region of interest, the lungs. Although we describe in detail the method for MCC measurement in mice, the protocol can be adjusted to study MCC in rats. The collimators would need to be adjusted as well as radiation dose. In our view, mouse MCC scans are more technically challenging due to the small animal size, but more useful than rats due to the large number of established mouse models of a number of human disorders. Additionally, due to their lower cost and cost of maintenance in animal colonies, a larger sample size is more feasible in mice.

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in vivo evaluation of mucociliary clearance in mice

Respiratory motile cilia, specialized organelles of the cell, line the apical surface of epithelial cells lining the respiratory tract. By beating in a metachronal, synchronal fashion, these multiple, motile, actin-based organelles generate a cephalad fluid flow clearing the respiratory tract of inhaled pollutants and pathogens. With increasing environmental pollution, novel viral pathogens and emerging multi-drug resistant bacteria, cilia generated mucociliary clearance (MCC) is essential for maintaining lung health. MCC is also depressed in multiple congenital disorders like primary ciliary dyskinesia, cystic fibrosis as well as acquired disorders like chronic obstructive pulmonary disease. All these disorders have established, in some case multiple, mouse models. In this publication, we detail a method using a small amount of radioactivity and dual-modality SPECT/CT imaging to accurately and reproducibly measure MCC in mice in vivo. The method allows for recovery of mice after imaging, making serial measurements possible, and testing potential therapeutics longitudinally over time. The data in wild-type mice demonstrates the reproducibility of the MCC measurement as long as adequate attention to detail is paid, and the protocol strictly adhered to.

Using this protocol, we anesthetized mice in an isoflurane chamber (Figure 1A). After achieving an adequate level of anesthesia, mice were placed onto vertical supports (Figure 1B) and the vocal cords were visualized using an illuminated guide wire (Figure 1C-1D). The mice were intubated and instilled with 0.2 mCi 99mTc-Sc in volumes of 10 &#181;L through a cannula and mice allowed to spontaneously inhale into the lungs (Figure 1E-1F). After image acquisition and processing, the CT and SPECT images were colocalized (Figure 2A) using the phantom tube as a landmark (Figure 2B). Masks of the lungs were generated from the CT image (Figure 2C) and used to draw ROIs around the right lung for analysis at 0 (Figure 2D) and 6 hours (Figure 2E-2F). To test reproducibility of the protocol, a total of 8 mice were scanned twice on different days with identical experimental conditions, with analysis using a paired t-test showing no significant difference between the repeat scans (p-value=0.9904) (Figure 3A). An additional 2 mice were scanned three times on different days with identical experimental conditions, with analysis using one-way ANOVA showing significant matching between the repeat scans (p-value of 0.0041) (Figure 3B). A total of 8 mice were scanned and two representative images were displayed (Figure 4).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/61929/61929fig01.jpg" /> 
Figure 1: Mouse intubation and isotope instillation.&#160;Images of the steps needed to intubate and instill isotope into the airway. A) The mouse is anesthetized in a chamber. B) The anesthetized mouse is placed onto a vertical support, suspended by the front incisors. C) An illuminated 0.5 mm fiber optic wire serving as a guide wire is prepared by running it through a 20 G cannula. D) The mouth of the mouse is opened using forceps and illuminated using the lighted guidewire to visualize the vocal cords. E) The cannula is pushed across the vocal cords and guidewire is removed. F) Soluble isotope is instilled into the cannula using a pipette and the mouse allowed to spontaneously inhale the isotope into the lungs. <a href="https://www.jove.com/files/ftp_upload/61929/61929fig01large.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/61929/61929fig02v2.jpg" /> 
Figure 2: SPECT/CT images of an MCC scan.&#160;A) A SPECT image that has been co-localized with a CT image. B) A CT image with a visible phantom tube that was used for co-localization. C) A mask of the airway derived by binarizing the CT image and performing a z-stack sum projection. D) The CT mask co-localized with the SPECT image. An ROI for analysis has been drawn around the right lung. E) A mask of the airway at 6 hours. F) A CT and SPECT co-localized image of the airway at 6 hours with an ROI for analysis. <a href="https://www.jove.com/files/ftp_upload/61929/61929fig02large.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/61929/61929fig03.jpg" /> 
Figure 3: Clearance measurements of the same mice across multiple scans.&#160;A) Two individual repeat clearance were measured for 8 mice with no changes in experimental conditions. A paired t test showed that there was no significant difference between the repeat scans with a p-value of 0.9904. B) Three individual repeat clearances were measured for two mice with no changes in experimental conditions. A one-way ANOVA showed there was significant matching between the repeat scans with a p-value of 0.0041. <a href="https://www.jove.com/files/ftp_upload/61929/61929fig03large.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/61929/61929fig04.jpg" /> 
Figure 4: Co-localized SPECT/CT images of the 0 and 6 hour airway in 2 mice with ROIs drawn at 0 and 6 hours outlining the right lung. <a href="https://www.jove.com/files/ftp_upload/61929/61929fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Supplemental Figure 1: A video of the vocal chords illuminated by a fiber optic wire with the effect of breathing visualized. <a href="https://cloudflare.jove.com/files/ftp_upload/61929/61929supfig01.jpg" target="_blank">Please click here to download this figure.</a>
Supplemental Files. <a href="https://cloudflare.jove.com/files/ftp_upload/61929/Supplemental_Files.zip" target="_blank">Please click here to download these files.</a>

Watch this Scientific Journal Video about In vivo Evaluation of Mucociliary Clearance in Mice at JoVE.com

In vivo Evaluation of Mucociliary Clearance in Mice

In this publication, we describe protocols for assessing airway mucociliary clearance (MCC) in mice in vivo utilizing dual-modality radionuclide imaging. This protocol is designed for a single photon emission computed tomography (SPECT) and computed tomography (CT) acquisition protocol using mouse whole body (MWB) collimators in a dual SPECT/CT system.

Respiratory motile cilia, specialized organelles of the cell, line the apical surface of epithelial cells lining the respiratory tract. By beating in a ...

in-vivo-evaluation-of-mucociliary-clearance-in-mice

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4.2K Views.  University of Pittsburgh School of Medicine. In this publication, we describe protocols for assessing airway mucociliary clearance (MCC) in mice in vivo utilizing dual-modality radionuclide imaging. This protocol is designed for a single photon emission computed tomography (SPECT) and computed tomography (CT) acquisition protocol using mouse whole body (MWB) collimators in a dual SPECT/CT system.

Video: In vivo Evaluation of Mucociliary Clearance in Mice

In Vivo 2-Photon Calcium Imaging in Layer 2/3 of Mice

To understand network dynamics of microcircuits in the neocortex, it is essential to simultaneously record the activity of a large number of neurons . In-vivo two-photon calcium imaging is the only method that allows one to record the activity of a dense neuronal population with single-cell resolution . The method consists in implanting a cranial imaging window, injecting a fluorescent calcium indicator dye that can be taken up by large numbers of neurons and finally recording the activity of neurons with time lapse calcium imaging using an in-vivo two photon microscope. Co-injection of astrocyte-specific dyes allows one to differentiate neurons from astrocytes. The technique can be performed in mice expressing fluorescent molecules in specific subpopulations of neurons to better understand the network interactions of different groups of cells.

To understand network dynamics of microcircuits in the neocortex, it is essential to simultaneously record the activity of a large number of neurons . In-vivo two-photon calcium imaging is the only method that allows one to record the activity of a dense neuronal population with single-cell resolution .

To understand network dynamics of microcircuits in the neocortex, it is essential to simultaneously record the activity of a large number of neurons .  ...

Evaluation of the Spatial Distribution of &gamma;H2AX following Ionizing Radiation

An early molecular response to DNA double-strand breaks (DSBs) is phosphorylation of the Ser-139 residue within the terminal SQEY motif of the histone H2AX1,2. This phosphorylation of H2AX is mediated by the phosphatidyl-inosito 3-kinase (PI3K) family of proteins, ataxia telangiectasia mutated (ATM), DNA-protein kinase catalytic subunit and ATM and RAD3-related (ATR)3. The phosphorylated form of H2AX, referred to as &gamma;H2AX, spreads to adjacent regions of chromatin from the site of the DSB, forming discrete foci, which are easily visualized by immunofluorecence microscopy3. Analysis and quantitation of &gamma;H2AX foci has been widely used to evaluate DSB formation and repair, particularly in response to ionizing radiation and for evaluating the efficacy of various radiation modifying compounds and cytotoxic compounds4.
Given the exquisite specificity and sensitivity of this de novo marker of DSBs, it has provided new insights into the processes of DNA damage and repair in the context of chromatin. For example, in radiation biology the central paradigm is that the nuclear DNA is the critical target with respect to radiation sensitivity. Indeed, the general consensus in the field has largely been to view chromatin as a homogeneous template for DNA damage and repair. However, with the use of &gamma;H2AX as molecular marker of DSBs, a disparity in &gamma;-irradiation-induced &gamma;H2AX foci formation in euchromatin and heterochromatin has been observed5-7. Recently, we used a panel of antibodies to either mono-, di- or tri- methylated histone H3 at lysine 9 (H3K9me1, H3K9me2, H3K9me3) which are epigenetic imprints of constitutive heterochromatin and transcriptional silencing and lysine 4 (H3K4me1, H3K4me2, H3K4me3), which are tightly correlated actively transcribing euchromatic regions, to investigate the spatial distribution of &gamma;H2AX following ionizing radiation8. In accordance with the prevailing ideas regarding chromatin biology, our findings indicated a close correlation between &gamma;H2AX formation and active transcription9. Here we demonstrate our immunofluorescence method for detection and quantitation of &gamma;H2AX foci in non-adherent cells, with a particular focus on co-localization with other epigenetic markers, image analysis and 3D-modeling.

Evaluation of the Spatial Distribution of &amp;#947;H2AX following Ionizing Radiation

Microscopic analysis of &gamma;H2AX foci, which form following the phosphorylation of H2AX at Ser-139 in response to DNA double-strand breaks, has become an invaluable tool in radiation biology. Here we used an antibody to mono-methylated histone H3 at lysine 4 as an epigenetic marker of actively transcribing euchromatin, to evaluate the spatial distribution of radiation-induced &gamma;H2AX formation within the nucleus.

An early molecular response to DNA double-strand breaks (DSBs) is phosphorylation of the Ser-139 residue within the terminal SQEY motif of the histone ...

Patch Clamp and Perfusion Techniques for Studying Ion Channels Expressed in Xenopus oocytes

The protocol presented here is designed to study the activation of the large conductance, voltage- and Ca2+-activated K+ (BK) channels. The protocol may also be used to study the structure-function relationship for other ion channels and neurotransmitter receptors1. BK channels are widely expressed in different tissues and have been implicated in many physiological functions, including regulation of smooth muscle contraction, frequency tuning of inner hair cells and regulation of neurotransmitter release2-6. BK channels are activated by membrane depolarization and by intracellular Ca2+ and Mg2+ 6-9. Therefore, the protocol is designed to control both the membrane voltage and the intracellular solution. In this protocol, messenger RNA of BK channels is injected into Xenopus laevis oocytes (stage V-VI) followed by 2-5 days of incubation at 18&deg;C10-13. Membrane patches that contain single or multiple BK channels are excised with the inside-out configuration using patch clamp techniques10-13. The intracellular side of the patch is perfused with desired solutions during recording so that the channel activation under different conditions can be examined. To summarize, the mRNA of BK channels is injected into Xenopus laevis oocytes to express channel proteins on the oocyte membrane; patch clamp techniques are used to record currents flowing through the channels under controlled voltage and intracellular solutions.

Patch Clamp and Perfusion Techniques for Studying Ion Channels Expressed in Xenopus oocytes

Ionic current of BK channels is recorded using patch clamp techniques. BK channels are expressed in Xenopus oocytes by injecting messenger RNA. The intracellular solution during patch clamp recordings is controlled by a perfusion system.

The protocol presented here is designed to study the activation of the large conductance, voltage- and Ca2+-activated K+ (BK) channels. The protocol may ...

Performing Vaginal Lavage, Crystal Violet Staining, and Vaginal Cytological Evaluation for Mouse Estrous Cycle Staging Identification

A rapid means of assessing reproductive status in rodents is useful not only in the study of reproductive dysfunction but is also required for the production of new mouse models of disease and investigations into the hormonal regulation of tissue degeneration (or regeneration) following pathological challenge. The murine reproductive (or estrous) cycle is divided into 4 stages: proestrus, estrus, metestrus, and diestrus. Defined fluctuations in circulating levels of the ovarian steroids 17-&beta;-estradiol and progesterone, the gonadotropins luteinizing and follicle stimulating hormones, and the luteotropic hormone prolactin signal transition through these reproductive stages. Changes in cell typology within the murine vaginal canal reflect these underlying endocrine events. Daily assessment of the relative ratio of nucleated epithelial cells, cornified squamous epithelial cells, and leukocytes present in vaginal smears can be used to identify murine estrous stages. The degree of invasiveness, however, employed in collecting these samples can alter reproductive status and elicit an inflammatory response that can confound cytological assessment of smears. Here, we describe a simple, non-invasive protocol that can be used to determine the stage of the estrous cycle of a female mouse without altering her reproductive cycle. We detail how to differentiate between the four stages of the estrous cycle by collection and analysis of predominant cell typology in vaginal smears and we show how these changes can be interpreted with respect to endocrine status.

Here, we describe how to identify the stage of the murine reproductive (proestrus, estrus, metestrus, or diestrus) by simple, non-invasive collection and cytological assessment of vaginal smear samples. We further describe how vaginal cytology reflects circulating hormonal levels underlying transition through the murine reproductive cycle.

A rapid means of assessing reproductive status in rodents is useful not only in the study of reproductive dysfunction but is also required for the ...

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

The proper elimination of unwanted or aberrant cells through apoptosis and subsequent phagocytosis (apoptotic cell clearance) is crucial for normal development in all metazoan organisms. Apoptotic cell clearance is a highly dynamic process intimately associated with cell death; unengulfed apoptotic cells are barely seen in vivo under normal conditions. In order to understand the different steps of apoptotic cell clearance and to compare 'professional' phagocytes - macrophages and dendritic cells to 'non-professional' - tissue-resident neighboring cells, in vivo live imaging of the process is extremely valuable. Here we describe a protocol for studying apoptotic cell clearance in live Drosophila embryos. To follow the dynamics of different steps in phagocytosis we use specific markers for apoptotic cells and phagocytes. In addition, we can monitor two phagocyte systems in parallel: 'professional' macrophages and 'semi-professional' glia in the developing central nervous system (CNS). The method described here employs the Drosophila embryo as an excellent model for real time studies of apoptotic cell clearance.

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

Here we describe an effective method for studying dynamics of apoptotic cell clearance in vivo. This method employs live Drosophila embryos as a powerful model for monitoring phagocytosis of apoptotic cells using specific labeling of apoptotic cells and phagocytes.

The  proper elimination of unwanted or aberrant cells through apoptosis and  subsequent phagocytosis (apoptotic cell clearance) is crucial for normal  ...

Evaluation of Respiratory System Mechanics in Mice using the Forced Oscillation Technique

The forced oscillation technique (FOT) is a powerful, integrative and translational tool permitting the experimental assessment of lung function in mice in a comprehensive, detailed, precise and reproducible manner. It provides measurements of respiratory system mechanics through the analysis of pressure and volume signals acquired in reaction to predefined, small amplitude, oscillatory airflow waveforms, which are typically applied at the subject's airway opening. The present protocol details the steps required to adequately execute forced oscillation measurements in mice using a computer-controlled piston ventilator (flexiVent; SCIREQ Inc, Montreal, Qc, Canada). The description is divided into four parts: preparatory steps, mechanical ventilation, lung function measurements, and data analysis. It also includes details of how to assess airway responsiveness to inhaled methacholine in anesthetized mice, a common application of this technique which also extends to other outcomes and various lung pathologies. Measurements obtained in na&iuml;ve mice as well as from an oxidative-stress driven model of airway damage are presented to illustrate how this tool can contribute to a better characterization and understanding of studied physiological changes or disease models as well as to applications in new research areas.

The present protocol provides a detailed step-by-step description of the procedures required to execute measurements of respiratory system mechanics as well as the assessment of airway responsiveness to inhaled methacholine in mice using the forced oscillation technique (flexiVent; SCIREQ Inc, Montreal, Qc, Canada).

The  forced oscillation technique (FOT) is a powerful, integrative and translational  tool permitting the experimental assessment of lung function in mice ...

Evaluation of Muscle Function of the Extensor Digitorum Longus Muscle Ex vivo and Tibialis Anterior Muscle In situ in Mice

Body movements are mainly provided by mechanical function of skeletal muscle. Skeletal muscle is composed of numerous bundles of myofibers that are sheathed by intramuscular connective tissues. Each myofiber contains many myofibrils that run longitudinally along the length of the myofiber. Myofibrils are the contractile apparatus of muscle and they are composed of repeated contractile units known as sarcomeres. A sarcomere unit contains actin and myosin filaments that are spaced by the Z discs and titin protein. Mechanical function of skeletal muscle is defined by the contractile and passive properties of muscle. The contractile properties are used to characterize the amount of force generated during muscle contraction, time of force generation and time of muscle relaxation. Any factor that affects muscle contraction (such as interaction between actin and myosin filaments, homeostasis of calcium, ATP/ADP ratio, etc.) influences the contractile properties. The passive properties refer to the elastic and viscous properties (stiffness and viscosity) of the muscle in the absence of contraction. These properties are determined by the extracellular and the intracellular structural components (such as titin) and connective tissues (mainly collagen) 1-2. The contractile and passive properties are two inseparable aspects of muscle function. For example, elbow flexion is accomplished by contraction of muscles in the anterior compartment of the upper arm and passive stretch of muscles in the posterior compartment of the upper arm. To truly understand muscle function, both contractile and passive properties should be studied.
The contractile and/or passive mechanical properties of muscle are often compromised in muscle diseases. A good example is Duchenne muscular dystrophy (DMD), a severe muscle wasting disease caused by dystrophin deficiency 3. Dystrophin is a cytoskeletal protein that stabilizes the muscle cell membrane (sarcolemma) during muscle contraction 4. In the absence of dystrophin, the sarcolemma is damaged by the shearing force generated during force transmission. This membrane tearing initiates a chain reaction which leads to muscle cell death and loss of contractile machinery. As a consequence, muscle force is reduced and dead myofibers are replaced by fibrotic tissues 5. This later change increases muscle stiffness 6. Accurate measurement of these changes provides important guide to evaluate disease progression and to determine therapeutic efficacy of novel gene/cell/pharmacological interventions. Here, we present two methods to evaluate both contractile and passive mechanical properties of the extensor digitorum longus (EDL) muscle and the contractile properties of the tibialis anterior (TA) muscle.

Evaluation of Muscle Function of the Extensor Digitorum Longus Muscle Ex vivo and Tibialis Anterior Muscle In situ in Mice

Changes in limb muscle contractile and passive mechanical properties are important biomarkers for muscle diseases. This manuscript describes physiological assays to measure these properties in the murine extensor digitorum longus and tibialis anterior muscles.

Body  movements are mainly provided by mechanical function of skeletal muscle. Skeletal  muscle is composed of numerous bundles of myofibers that are ...

In Vivo SiRNA Transfection and Gene Knockdown in Spinal Cord via Rapid Noninvasive Lumbar Intrathecal Injections in Mice

This report describes a step-by-step guide to the technique of acute intrathecal needle injections in a noninvasive manner, i.e. independent of catheter implantation. The technical limitation of this surgical technique lies in the finesse of the hands. The injection is rapid, especially for a trained experimenter, and since tissue disruption with this technique is minimal, repeated injections are possible; moreover immune reaction to foreign tools (e.g. catheter) does not occur, thereby giving a better and more specific read out of spinal cord modulation. Since the application of the substance is largely limited to the target region of the spinal cord, drugs do not need to be applied in large dosages, and more importantly unwanted effects on other tissue, as observed with a systemic delivery, could be circumvented1,2. Moreover, we combine this technique with in vivo transfection of nucleic acid with the help of polyethylenimine (PEI) reagent3, which provides tremendous versatility for studying spinal functions via delivery of pharmacological agents as well as gene, RNA, and protein modulators.

In Vivo SiRNA Transfection and Gene Knockdown in Spinal Cord via Rapid Noninvasive Lumbar Intrathecal Injections in Mice

This report describes a simple and rapid technique of intrathecal needle puncture for a localized transfection of siRNA in the lumbar spinal cord in mouse under short lasting light anesthesia.

This report describes a step-by-step guide to the technique of acute intrathecal needle injections in a noninvasive manner, i.e. independent of catheter ...

Evaluation of Zebrafish Kidney Function Using a Fluorescent Clearance Assay

The zebrafish embryo offers a tractable model to study organogenesis and model human genetic disease. Despite its relative simplicity, the zebrafish kidney develops and functions in almost the same way as humans. A major difference in the construction of the human kidney is the presence of millions of nephrons compared to the zebrafish that has only two. However, simplifying such a complex system into basic functional units has aided our understanding of how the kidney develops and operates. In zebrafish, the midline located glomerulus is responsible for the initial blood filtration into two pronephric tubules that diverge to run bilaterally down the embryonic axis before fusing to each other at the cloaca. The pronephric tubules are heavily populated by motile cilia that facilitate the movement of filtrate along the segmented tubule, allowing the exchange of various solutes before finally exiting via the cloaca2-4. Many genes responsible for CKD, including those related to ciliogenesis, have been studied in zebrafish5. However, a major draw back has been the difficulty in evaluating zebrafish kidney function after genetic manipulation. Traditional assays to measure kidney dysfunction in humans have proved non translational to zebrafish, mainly due to their aquatic environment and small size. For example, it is not physically possible to extract blood from embryonic staged fish for analysis of urea and creatinine content, as they are too small. In addition, zebrafish do not produce enough urine for testing on a simple proteinuria &#8216;dipstick&#8217;, which is often performed during initial patient examinations. We describe a fluorescent assay that utilizes the optical transparency of the zebrafish to quantitatively monitor the clearance of a fluorescent dye, over time, from the vasculature and out through the kidney, to give a read out of renal function1,6-9.

The zebrafish is a popular tool to model chronic kidney disease (CKD). However, their small size makes it impossible to evaluate renal function using traditional methods. We describe a fluorescent dye kidney clearance assay1 that allows quantitative analysis of zebrafish kidney function in CKD.

The zebrafish embryo offers a tractable model to study organogenesis and model human genetic disease. Despite its relative simplicity, the zebrafish ...

Measuring the Stiffness of Ex Vivo Mouse Aortas Using Atomic Force Microscopy

Arterial stiffening is a significant risk factor and biomarker for cardiovascular disease and a hallmark of aging. Atomic force microscopy (AFM) is a versatile analytical tool for characterizing viscoelastic mechanical properties for a variety of materials ranging from hard (plastic, glass, metal, etc.) surfaces to cells on any substrate. It has been widely used to measure the stiffness of cells, but less frequently used to measure the stiffness of aortas. In this paper, we will describe the procedures for using AFM in contact mode to measure the ex vivo elastic modulus of unloaded mouse arteries. We describe our procedure for isolation of mouse aortas, and then provide detailed information for the AFM analysis. This includes step-by-step instructions for alignment of the laser beam, calibration of the spring constant and deflection sensitivity of the AFM probe, and acquisition of force curves. We also provide a detailed protocol for data analysis of the force curves.

Measuring the Stiffness of Ex Vivo Mouse Aortas Using Atomic Force Microscopy

We present detailed protocols for isolation of aortas from mouse and measurement of their elastic modulus using atomic force microscopy.

Arterial stiffening is a significant risk factor and biomarker for cardiovascular disease and a hallmark of aging. Atomic force microscopy (AFM) is a ...

Methods Article

In vivo Evaluation of Mucociliary Clearance in Mice