We thank Dr. Harvey Hensley for his expert technical support.  This work benefited from the use of the following Fox Chase Cancer Center (FCCC) facilities: Laboratory of Animal Facility and Small Animal Imaging Facility.  This work was supported in part by the Ovarian Cancer SPORE at FCCC/UPenn (P50 CA083638), the FCCC core grant (P30 CA006927), and the Ovarian Cancer Research Fund.

I. Preparation of Ovarian Tumor Cells
<ol>
<li>	Grow ovarian cancer cell lines expressing luciferase in culture. Sources of luciferase can be Firefly, Renilla, or other species. Cells expressing fluorescent proteins can also be used. For this demonstration we use an ovarian cancer cell line, OVCAR5, expressing firefly luciferase.</li>
<li>	Harvest cells using routine cell culture technique.</li>
<li>	Keep the cell suspension (10,000 cells/&mu;L) in phosphate buffered saline (PBS) on ice until time of injection.</li>
</ol>

II. Intrabursal Injection
This procedure requires assistance from a second person. All surgical procedures are conducted under aseptic conditions. This includes wearing surgical attire and using sterile surgical instruments, syringe, and needles.
<ol>
<li>	Prepare the anesthetic solution by mixing 3 mL of ketamine hydrochloride (100 mg/mL), 1.6 mL of xylazine hydrochloride (100 mg/mL), 1.5 mL of acepromazine (10 mg/mL), and 20 mL of 0.9% sodium chloride.</li>
<li>	Check the animal's identification number and observable health. Anesthetize the animal with prepared ketamine-xylazine-acepromazine anesthesia via intraperitoneal (i.p.) injection with a dose volume of 8~9 mL/kg body weight (BW).</li>
<li>	Confirm that the animal is under an acceptable plain of anesthesia by performing a toe pinch with forceps or fingers to the animal's hind paws. If there is pedal reflex, wait for a deeper plain of anesthesia until the animal is unresponsive to this procedure.</li>
<li>	Lay the animal dorsal side up on a sterile gauze pad with its head facing away and its tail facing towards you. The point of incision is located to the left or right of the midline and above the ovaries. Shave or wet the fur with 70% alcohol at the incision point.</li>
<li>	Lift the wetted skin using forceps and make a small incision with the scissor at the dorsomedial position and directly above the ovarian fat pad. The ovarian fat pad should be visible beneath the surface of the peritoneal wall. Fat pad is easily recognizable by its white color in contrast to the dark pink tissue surrounding it.</li>
<li>	Gently lift the peritoneal wall lining and make a small incision as described above (5).</li>
<li>	Place a sterile soaked saline gauze pad on the midline adjacent to the incision. Locate the ovarian fat pad and gently pull it out and rest it onto the gauze. Stabilize the ovary by clamping the fat pad with a bulldog clip. Under a dissecting microscope, position the ovary as to allow for the insertion of the needle (30 gauge, G) into the oviduct tubule bend leading to the bursa. When the needle is inserted into the proper position, it should be visible under the bursa.</li>
<li>	Gently push the plunger of the syringe to inject 5 &mu;L of cell suspension between the bursa and the ovary while the syringe is positioned to injection site. This step requires two people. One person pushes the plunger while the other person maintains positioning of needle. Remove the needle quickly to seal the puncture site but gently enough not to tear the bursa and tubule. The bursa should appear to be slightly distended with proper injection.</li>
<li>	Release the fat pad from the bulldog clip and gently replace the reproductive tract and fat pad back into the peritoneal cavity. Gently close the body wall by pulling the upper peritoneal lining over the lower lining. Close the skin with surgical staples or wound clips.</li>
<li>	Place the recovering animal back in its cage and provide a safe heat source to avoid hypothermia and speed up recovery. Monitor the breathing rate and ease, the return of muscle tone, and the ability to voluntarily move. These are all good indicators of the progression towards recovery. Staples or wound clips can be removed 7 or more days post surgery.</li>
</ol>

III. Oral Gavage Administration
<ol>
<li>	Use an 18~20 G gavage needle or feeding tube with a rounded tip. The gavage needle should be no longer than the distance from the tip of the animal's head to its last rib.</li>
<li>	Check the animals' identification number and observable health. Gently scruff the animal by grasping the skin over the shoulders with thumb and fingers. The restraint should be just firm enough for the fore limbs to be extended to each side and out of the way. The animal should not be able to grasp at the needle.</li>
<li>	Gently pull back the animal's head with your index finger, forming a straight line through the neck and esophagus. Support the animal's back with the inside of your thumb.</li>
<li>	Gently and without force, insert the gavage needle at either side of the mouth and over the tongue. In one smooth motion, the needle should pass against the roof of the mouth and down the esophagus. Gravity should guide the needle down without meeting resistance. If there is any resistance, remove the needle and start again. The animal's gag and swallow reflex may be triggered during insertion. However, there should be no resistance or gasping from the animal. It is important to make sure that the animal is breathing when the needle is in place by checking the movement of the nostrils and chest.</li>
<li>	Slowly push plunger of the syringe to dispense the dose volume.</li>
<li>	Gently remove the gavage needle at the same angle and pathway as the way it was inserted down the esophagus.</li>
<li>	Return the animal to its cage and monitor breathing and behavior for 5~10 minutes.</li>
</ol>

IV. In Vivo Imaging
We use the Caliper Life Sciences to monitor the behavior of cells injected into intrabursal cavity. Experiments using this system typically have a timeframe of 4~16 weeks from the time of tumor implantation.
<ol>
<li>	Prepare the solution of D-luciferin (substrate of firefly luciferase) by dissolving 5~20 mg in 1 mL PBS and filtering through 0.22 &mu;m membrane for sterilization.</li>
<li>	Charge the induction chamber by turning on both the oxygen and the isoflurane gas. Turn ON the gas flow to the induction chamber only. The isoflurane flow rate is maintained at a low level until animals are ready to be imaged.</li>
<li>	Check the animals' identification number and observable health. Place the animal in the induction chamber charged with isoflurane gas.</li>
<li>	Remove the animal from the chamber when the animal appears to be anesthetized. Deliver 200 &mu;L of prepared luciferin solution via i.p. injection using 30 G needle. It is important to record the time of injection in order to keep the time consistent between luciferin injection and imaging performance throughout the study.</li>
<li>	Place the luciferin-injected animal back into the induction chamber.</li>
<li>	Set the imaging system to the appropriate settings. Be sure to initialize the system before acquiring the first animal image.</li>
<li>	Turn OFF the gas flow to the induction chamber and turn ON the gas flow to the imaging chamber.</li>
<li>	Place the anesthetized animal dorsal or ventral side up. Be sure to place the animals' nose in a nose cone to maintain proper anesthesia induction. Make sure the animal is within the "point of interest grid" as indicated by green illumination.</li>
<li>	Close the door of the chamber and acquire the image at the appropriate time. Time can be determined by kinetics.</li>
<li>	Remove the animal from the imaging chamber and place it back into its cage. Monitor the animals' recovery as described in "Intrabursal Injection".</li>
</ol>

V. Representative Results
<img src="/files/ftp_upload/2125/2125fig1.jpg" alt="Figure 1" /> Figure 1. In Vivo imaging of ovarian tumor cells in an orthotopic mouse model. OVCAR5 cells expressing luciferase were injected intrabursally into right ovary and imaged over time. Images were taken 13 (left), 17 (middle), and 22 (right) days after the injection using the IVIS Spectrum imaging system. Dorsal side is shown in upper panel and ventral side is in lower panel. Note that the peritoneal spread of tumor cells 22 days following injection.
Experiments on animals were performed in accordance with the guidelines and regulations set forth by Fox Chase Cancer Center's Institutional Animal Care and Use Committee.

<ol>
	<li>Jemal, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cancer+statistics.">Cancer statistics.</a> CA Cancer J Clin. 59, 225-249 (2009).</li><li>Lynch, H. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Hereditary+ovarian+carcinoma:+heterogeneity,+molecular+genetics,+pathology,+and+management.">Hereditary ovarian carcinoma: heterogeneity, molecular genetics, pathology, and management.</a> Mol Oncol. 3, 97-137 (2009).</li><li>Yap, T. A., Carden, C. P., Kaye, S. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Beyond+chemotherapy:+targeted+therapies+in+ovarian+cancer.">Beyond chemotherapy: targeted therapies in ovarian cancer.</a> Nat Rev Cancer. 9, 167-181 (2009).</li><li>Killion, J. J., Radinsky, R., Fidler, I. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Orthotopic+models+are+necessary+to+predict+therapy+of+transplantable+tumors+in+mice.">Orthotopic models are necessary to predict therapy of transplantable tumors in mice.</a> Cancer Metastasis Rev. 17, 279-284 (1998).</li><li>Bibby, M. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Orthotopic+models+of+cancer+for+preclinical+drug+evaluation:+advantages+and+disadvantages.">Orthotopic models of cancer for preclinical drug evaluation: advantages and disadvantages.</a> Eur J Cancer. 40, 852-857 (2004).</li><li>Shaw, T. J., Senterman, M. K., Dawson, K., Crane, C. A., Vanderhyden, B. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Characterization+of+intraperitoneal,+orthotopic,+and+metastatic+xenograft+models+of+human+ovarian+cancer.">Characterization of intraperitoneal, orthotopic, and metastatic xenograft models of human ovarian cancer.</a> Mol Ther. 10, 1032-1042 (2004).</li><li>Greenaway, J., Moorehead, R., Shaw, P., Petrik, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Epithelial-stromal+interaction+increases+cell+proliferation,+survival+and+tumorigenicity+in+a+mouse+model+of+human+epithelial+ovarian+cancer.">Epithelial-stromal interaction increases cell proliferation, survival and tumorigenicity in a mouse model of human epithelial ovarian cancer.</a> Gynecol Oncol. 108, 385-394 (2008).</li><li>Connolly, D. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Animal+models+of+ovarian+cancer.">Animal models of ovarian cancer.</a> Cancer Treat Res. 149, 353-391 (2009).</li><li>Sadikot, R. T., Blackwell, T. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Bioluminescence+imaging.">Bioluminescence imaging.</a> Proc Am Thorac Soc. 2, 537-540 (2005).</li><li>Lehmann, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Longitudinal+and+multimodal+in+vivo+imaging+of+tumor+hypoxia+and+its+downstream+molecular+events.">Longitudinal and multimodal in vivo imaging of tumor hypoxia and its downstream molecular events.</a> Proc Natl Acad Sci U S A. 106, 14004-14009 (2009).</li><li>Connolly, D. C., Hensley, H. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Xenograft+and+Transgenic+Mouse+Models+of+Epithelial+Ovarian+Cancer+and+Non-Invasive+Imaging+Modalities+to+Monitor+Ovarian+Tumor+Growth+In+Situ:+Applications+in+Evaluating+Novel+Therapeutic+Agents.">Xenograft and Transgenic Mouse Models of Epithelial Ovarian Cancer and Non-Invasive Imaging Modalities to Monitor Ovarian Tumor Growth In Situ: Applications in Evaluating Novel Therapeutic Agents.</a> Current Protocols in Pharmacoogy. 45, (2009).</li></ol>

No conflicts of interest declared.

Ovarian cancer is the leading cause of death among all gynecologic malignancies 1. The high mortality rate of this disease is largely due to its late diagnosis and the lack of reliable diagnostic methods 2. Furthermore, conventional chemotherapy often encounters chemoresistance and relapse of cancer 3. Therefore, novel therapeutics is required to effectively target this disease. In development of new therapy targeting human ovarian cancer, it is critical to develop a representative animal model.

An orthotopic animal model has advantages over conventional xenograft models (e.g. subcutaneous or intraperitoneal injections of tumor cells) in that 1) it reproduces the primary site of tumor formation, 2) it represents common site of metastases, and 3) it provides tumor cells to interact with appropriate microenvironment 4, 5, 6, 7, 8. Rodents have a unique bursal membrane that surrounds the ovary and is continuous with the oviduct. This unique anatomy of rodents allows injection of ovarian tumor cells orthotopically. Intrabursally injected ovarian tumor cells behave similar to human disease, thereby growing within intrabursal membrane and spreading into peritoneal cavity as tumor progresses. Injection of cells stably expressing luciferases or fluorescent proteins also allows tracking behavior of tumor cells in real time. Bioluminescent and/or fluorescent imaging technology makes it possible to repeatedly image tumor cells over an extended period of time and study tumor growth, distribution, and regression in non-invasive manner 9, 10, 11.

In establishing the orthotopic model, it is critical to quickly remove the needle from the bursa upon injection. Abrupt removal of the needle seals the puncture site and prevents the leakage of injected cells. However, at the same time, the movement should be gentle not to tear the bursa. If leakage occurs during injection, it can cause cells to seed in the abdomen and potentially confound the study, i.e., premature spreading outside the ovary. In this event, the specific animal should be recorded and followed for the development of multiple tumor sites in the peritoneum prior to treatment or alternatively eliminated from the subsequent analyses. Prior to performing oral gavage, it is critical to check the length of the gavage tube by measuring from the tip of the animal's head to the last rib. It is helpful to mark the tube at the animal's nose and do not pass the tube past this mark. Insertion of the tube past this mark can result in a perforation to the stomach. Determining the length of the tube is especially important with younger animals or animals weighing under 20 g. Upon insertion of the gavage needle down the esophagus, there should be no resistance or struggle from the animal. It is important not to administer the solution or suspension too fast as this can lead to reflux and subsequently deliver inaccurate dose volume, as well as adding stress to the animal. For comparable imaging results from time to time, it is desirable to keep the time consistent between injection of luciferase substrate and imaging. The time lapse can be determined by taking series of images after substrate injection and observe the kinetics of signals. Development of an orthotopic mouse model of ovarian cancer, oral delivery of drugs, and in vivo imaging are necessary techniques for better understanding of development and spread of ovarian cancer and assessing novel therapeutic regimens that may ultimately improve the outcome of patient with this deadly disease.

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in vivo imaging and therapeutic treatments in an orthotopic mouse model of ovarian cancer

Human cancer and response to therapy is better represented in orthotopic animal models. This paper describes the development of an orthotopic mouse model of ovarian cancer, treatment of cancer via oral delivery of drugs, and monitoring of tumor cell behavior in response to drug treatment in real time using in vivo imaging system. In this orthotopic model, ovarian tumor cells expressing luciferase are applied topically by injecting them directly into the mouse bursa where each ovary is enclosed. Upon injection of D-luciferin, a substrate of firefly luciferase, luciferase-expressing cells generate bioluminescence signals. This signal is detected by the in vivo imaging system and allows for a non-invasive means of monitoring tumor growth, distribution, and regression in individual animals. Drug administration via oral gavage allows for a maximum dosing volume of 10 mL/kg body weight to be delivered directly to the stomach and closely resembles delivery of drugs in clinical treatments. Therefore, techniques described here, development of an orthotopic mouse model of ovarian cancer, oral delivery of drugs, and in vivo imaging, are useful for better understanding of human ovarian cancer and treatment and will improve targeting this disease.

Watch this Scientific Journal Video about In vivo Imaging and Therapeutic Treatments in an Orthotopic Mouse Model of Ovarian Cancer at JoVE.com

In vivo Imaging and Therapeutic Treatments in an Orthotopic Mouse Model of Ovarian Cancer

Orthotopic animal models of ovarian cancer replicate better human disease and therefore enhance our understanding of cancer progression and tumor response to therapy. A mouse model receives an intrabursal injection of luciferase-expressing ovarian tumor cells. Treatment is administered via oral gavage. Tumor growth is monitored by in vivo imaging system.

In vivo Imaging and Therapeutic Treatments in an Orthotopic Mouse Model of Ovarian Cancer

Human cancer and response to therapy is better represented in orthotopic animal models.  This paper describes the development of an orthotopic mouse model ...

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Video: In vivo Imaging and Therapeutic Treatments in an Orthotopic Mouse Model of Ovarian Cancer

Orthotopic Mouse Model of Colorectal Cancer

The traditional subcutaneous tumor model is less than ideal for studying colorectal cancer. Orthotopic mouse models of colorectal cancer, which feature cancer cells growing in their natural location, replicate human disease with high fidelity. Two techniques can be used to establish this model. Both techniques are similar and require mouse anesthesia and laparotomy for exposure of the cecum. One technique involves injection of a colorectal cancer cell suspension into the cecal wall. Cancer cells are first grown in culture, harvested when subconfluent and prepared as a single cell suspension. A small volume of cells is injected slowly to avoid leakage. The other technique involves transplantation of a piece of subcutaneous tumor onto the cecum. A mouse with a previously established subcutaneous colorectal tumor is euthanized and the tumor is removed using sterile technique. The tumor piece is divided into small pieces for transplantation to another mouse. Prior to transplantation, the cecal wall is lightly damaged to facilitate tumor cell infiltration. The time to developing primary tumors and liver metastases will vary depending on the technique, cell line, and mouse species used. This orthotopic mouse model is useful for studying the natural progression of colorectal cancer and testing new therapeutic agents against colorectal cancer.

Two techniques can be used to establish this model: injection of a cancer cell suspension into the cecal wall or transplantation of a piece of subcutaneous tumor onto the cecum. This model is useful for studying the natural progression of colorectal cancer and testing new therapeutic agents against colorectal cancer.

The traditional subcutaneous tumor model is less than ideal for studying colorectal cancer. Orthotopic mouse models of colorectal cancer, which feature ...

Imaging In-Stent Restenosis: An Inexpensive, Reliable, and Rapid Preclinical Model

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In this video, we present the full procedure and pit-falls of a rat stent model suitable for high throughput stent research. We will show the surgical procedure of stent deployment, and the assessment of in-stent restenosis using the most elegant technique of OCT (Optical Coherence Tomography). This technique provides high accuracy in assessing plaque CSAs (cross section areas) and correlates well with histological sections, which require special and time consuming embedding and sectioning techniques. OCT imaging further allows longitudinal monitoring of the development of in-stent restenosis within the same animal compared to one-time snapshots using histology.

This video demonstrates how to use a preclinical inexpensive and reliable model to study pathobiological and pathophysiological processes of in-stent restenosis development. Longitudinal in vivo monitoring using OCT (Optical Coherence Tomography) and analysis of OCT images are also demonstrated.

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Tracking of HSPCs in relation to GFP positive osteoblastic cells is shown as an exemplary application of this method. This technique can be utilized to track any appropriately labeled hematopoietic or stromal cell of interest within the mouse calvarium bone marrow space.

In Vivo 4-Dimensional Tracking of Hematopoietic Stem and Progenitor Cells in Adult Mouse Calvarial Bone Marrow

The nature of the interactions between hematopoietic stem and progenitor cells (HSPCs) and bone marrow niches is poorly understood. Custom hardware modifications and a multi-step acquisition protocol allow the use of two-photon and confocal microscopy to image ex vivo labeled HSPCs homed within bone marrow areas, tracking interactions and movement.

Through a delicate balance between quiescence and proliferation, self renewal and production of differentiated progeny, hematopoietic stem cells (HSCs) ...

Validation of a Mouse Model to Disrupt LINC Complexes in a Cell-specific Manner

Nuclear migration and anchorage within developing and adult tissues relies heavily upon large macromolecular protein assemblies called LInkers of the Nucleoskeleton and Cytoskeleton (LINC complexes). These protein scaffolds span the nuclear envelope and connect the interior of the nucleus to components of the surrounding cytoplasmic cytoskeleton. LINC complexes consist of two evolutionary-conserved protein families, Sun proteins and Nesprins that harbor C-terminal molecular signature motifs called the SUN and KASH domains, respectively. Sun proteins are transmembrane proteins of the inner nuclear membrane whose N-terminal nucleoplasmic domain interacts with the nuclear lamina while their C-terminal SUN domains protrudes into the perinuclear space and interacts with the KASH domain of Nesprins. Canonical Nesprin isoforms have a variable sized N-terminus that projects into the cytoplasm and interacts with components of the cytoskeleton. This protocol describes the validation of a dominant-negative transgenic mouse strategy that disrupts endogenous SUN/KASH interactions in a cell-type specific manner. Our approach is based on the Cre/Lox system that bypasses many drawbacks such as perinatal lethality and cell nonautonomous phenotypes that are associated with germline models of LINC complex inactivation. For this reason, this model provides a useful tool to understand the role of LINC complexes during development and homeostasis in a wide array of tissues.

Nuclear envelope proteins play a central role in many basic biological processes and have been implicated in a variety of human diseases. This protocol describes a new Cre/Lox-based mouse model that allows for the spatiotemporal control of LINC complexes disruption.

Nuclear migration and anchorage within developing and adult tissues relies heavily upon large macromolecular protein assemblies called LInkers of the ...

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 ...

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Presented here is a protocol for the fabrication of a spheroid imaging device. This device enables dynamic or longitudinal fluorescence imaging of cancer cell spheroids. The protocol also offers a simple image processing procedure for the analysis of cancer cell invasion.

The invasion of cancer cells from the primary tumor into the adjacent healthy tissues is an early step in metastasis. Invasive cancer cells pose a major ...

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Platelets are produced by megakaryocytes, specialized cells located in the bone marrow. The possibility to image megakaryocytes in real time and their native environment was described more than 10 years ago and sheds new light on the process of platelet formation. Megakaryocytes extend elongated protrusions, called proplatelets, through the endothelial lining of sinusoid vessels. This paper presents a protocol to simultaneously image in real time fluorescently labeled megakaryocytes in the skull bone marrow and sinusoid vessels. This technique relies on a minor surgery that keeps the skull intact to limit inflammatory reactions. The mouse head is immobilized with a ring glued to the skull to prevent movements from breathing.
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In Vivo Two-photon Imaging of Megakaryocytes and Proplatelets in the Mouse Skull Bone Marrow

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Platelets are produced by megakaryocytes, specialized cells located in the bone marrow. The possibility to image megakaryocytes in real time and their ...

Imaging Neutrophil Migration in the Mouse Skin to Investigate Subcellular Membrane Remodeling Under Physiological Conditions

The study of immune cell recruitment and function in tissues has been a very active field over the last two decades. Neutrophils are among the first immune cells to reach the site of inflammation and to participate in the innate immune response during infection or tissue damage. So far, neutrophil migration has been successfully visualized using various in vitro experimental systems based on uniform stimulation, or confined migration under agarose, or micro-fluidic channels. However, these models do not recapitulate the complex microenvironment that neutrophils encounter in vivo. The development of multiphoton microscopy (MPM)-based techniques, such as intravital subcellular microscopy (ISMic), offer a unique tool to visualize and investigate neutrophil dynamics at subcellular resolutions under physiological conditions. In particular, the ear of a live anesthetized mouse provides an experimental advantage to follow neutrophil interstitial migration in real-time due to its ease of accessibility and lack of surgical exposure. ISMic provides the optical resolution, speed, and depth of acquisition necessary to track both cellular and, more importantly, subcellular processes in 3D over time (4D). Moreover, multi-modal imaging of the interstitial microenvironment (i.e., blood vessels, resident cells, extracellular matrix) can be readily accomplished using a combination of transgenic mice expressing select fluorescent markers, exogenous labeling via fluorescent probes, tissue intrinsic fluorescence, and second/third harmonic generated signals. This protocol describes 1) the preparation of neutrophils for adoptive transfer into the mouse ear, 2) different settings for optimal sub-cellular imaging, 3) strategies to minimize motion artifacts while maintaining a physiological response, 4) examples of membrane remodeling observed in neutrophils using ISMic, and 5) a workflow for the quantitative analysis of membrane remodeling in migrating neutrophils in vivo.

Neutrophil migration relies on the rapid and continuous remodeling of the plasma membrane in response to chemoattractant and its interactions with the extracellular microenvironment. Described herein is a procedure based on Intravital Subcellular Microscopy to investigate the dynamic of membrane remodeling in neutrophils injected in the ear of anesthetized mice.

The study of immune cell recruitment and function in tissues has been a very active field over the last two decades. Neutrophils are among the first ...

Imaging of Mitochondrial Peroxide in Living Yeast Cells Using mtHyper7

Imaging of mtHyPer7, a Ratiometric Biosensor for Mitochondrial Peroxide, in Living Yeast Cells

Mitochondrial dysfunction, or functional alteration, is found in many diseases and conditions, including neurodegenerative and musculoskeletal disorders, cancer, and normal aging. Here, an approach is described to assess mitochondrial function in living yeast cells at cellular and subcellular resolutions using a genetically encoded, minimally invasive, ratiometric biosensor. The biosensor, mitochondria-targeted HyPer7 (mtHyPer7), detects hydrogen peroxide (H2O2) in mitochondria. It consists of a mitochondrial signal sequence fused to a circularly permuted fluorescent protein and the H2O2-responsive domain of a bacterial OxyR protein. The biosensor is generated and integrated into the yeast genome using a CRISPR-Cas9 marker-free system, for&#160;more consistent expression compared to plasmid-borne constructs.
mtHyPer7 is quantitatively targeted to mitochondria, has no detectable effect on yeast growth rate or mitochondrial morphology, and provides a quantitative readout for mitochondrial H2O2 under normal growth conditions and upon exposure to oxidative stress. This protocol explains how to optimize imaging conditions using a spinning-disk confocal microscope system and perform quantitative analysis using freely available software. These tools make it possible to collect rich spatiotemporal information on mitochondria both within cells and among cells in a population. Moreover, the workflow described here can be used to validate other biosensors.

Author Spotlight: Advancing Mitochondrial Research - mtHyper7 Biosensor for Subcellular Analysis 

Hydrogen peroxide (H2O2) is both a source of oxidative damage and a signaling molecule. This protocol describes how to measure mitochondrial H2O2 using mitochondria-targeted HyPer7 (mtHyPer7), a genetically encoded ratiometric biosensor, in living yeast. It details how to optimize imaging conditions and perform quantitative cellular and subcellular analysis using freely available software.

Mitochondrial dysfunction, or functional alteration, is found in many diseases and conditions, including neurodegenerative and musculoskeletal disorders, ...

Quantification of Skeletal Muscle Fatty Infiltration via Decellularization

Decellularization-Based Quantification of Skeletal Muscle Fatty Infiltration

Fatty infiltration is the accumulation of adipocytes between myofibers in skeletal muscle and is a prominent feature of many myopathies, metabolic disorders, and dystrophies. Clinically in human populations, fatty infiltration is assessed using noninvasive methods, including computed tomography (CT), magnetic resonance imaging (MRI), and ultrasound (US). Although some studies have used CT or MRI to quantify fatty infiltration in mouse muscle, costs and insufficient spatial resolution remain challenging. Other small animal methods utilize histology to visualize individual adipocytes; however, this methodology suffers from sampling bias in heterogeneous pathology. This protocol describes the methodology to qualitatively view and quantitatively measure fatty infiltration comprehensively throughout intact mouse muscle and at the level of individual adipocytes using decellularization. The protocol is not limited to specific muscles or specific species and can be extended to human biopsy. Additionally, gross qualitative and quantitative assessments can be made with standard laboratory equipment for little cost, making this procedure more accessible across research laboratories.

Author Spotlight: Decellularization-Based Quantification of Skeletal Muscle Fatty Infiltration  

The present study describes decellularization-based methodologies for visualizing and quantifying intramuscular adipose tissue (IMAT) deposition through intact muscle volume, as well as quantifying metrics of individual adipocytes that comprise IMAT.