We thank Dr. W. Mintzer for the suggestion to perform fiberoptic orotracheal intubation. Supported by grants U19AI070973, R01AI057696, K02HL75243, and R01HL082487 from the National Institutes of Health.

Allergen challenge:
<ol>
	<li>C57BL/6 mice, 4-8 weeks of age, are anesthetized in an airtight plexiglass chamber purged with a 3.2% isoflurane in oxygen vapor mixture for 10 minutes to achieve deep general anesthesia.</li>
	<li>Intranasal allergen challenges (45&mu;L OVA (22.5 &mu;g) and 7&mu;L A. Oryzae (7 &mu;g), in PBS) are administered, every Tuesday, Thursday and Sunday, for a total of seven consecutive applications.</li>
</ol>
Anesthesia:
<ol start="3">
	<li>Prior to each allergen challenge, and following the 7th challenge, mice are administered an intraperitoneal injection of 48 mg/kg etomidate (2 mg/ml), prior to placement in a light-excluding receptacle.</li>
	<li>Subject remains in receptacle until a lack of observable neurological response is detected upon application of pressure to hind paws (5-10 min).</li>
 </ol>
Intubation:
<ol start="5">
	<li>A radiant heat lamp, maintained at a distance to ensure maintenance of ~37&deg;C body core temperature, should be focused on the subject throughout the procedure to prevent hypothermia. A rectal thermometer should be used at least initially to confirm euthermia regardless of the heat source used. CRITICAL STEPS All fluids and instruments received by experimental animals should be sterile; procedures should be performed under strict aseptic conditions. Prolonged hypothermia while under anesthesia will lead to aberrant data and/or death of animals. Competence with all invasive procedures should be developed using cadaveric animals before attempting work on live animals. Ophthalmic lubricant should be used to prevent corneal abrasions of animals under general anesthesia.</li>
	<li>Anesthetized mice are removed from receptacle and placed in the recumbent position (ventral side up), on plethysmograph table, adjusted to a 45&deg; angle.</li>
	<li>A rubber band encircling the table is inserted behind the top row of incisors so to secure the subject into place. With tweezers in right hand, grip, extend, and lift the tongue from the mouth before securing it in place with a metal depressor in left hand; thus allowing an unimpeded airway for intubation.</li>
	<li>A 0.8 mm diameter fiberoptic thread, connected to a light source, is inserted through the angiocatheter and extended 10 mm beyond the tip. As the depressor is steadied with left hand, the illuminated end of the fiberoptic thread is guided through the oral cavity and pharynx by the right hand until the vocal chords are visualized. The thread is then passed under direct visualization through the moving vocal cords and into the trachea, timed to occur when the cords are maximally open.</li>
	<li>The angiocatheter is then passed over the fiberoptic thread into the trachea until the catheter tip lies within the mid-portion of the trachea. For 17-22 gram mice, this corresponds to a 10 mm catheter segment remaining visible between the connector and the cranial extreme of the subject s lower jaw. The actual amount by which the catheter is passed should be determined by direct inspection of the trachea of 2-3 catheterized mice of the relevant size and genetic background.</li>
	<li>The fiberoptic thread is removed and successful intubation is confirmed by observing regular deep breaths (rhythmic excursions of the thorax and abdomen) which immediately terminate following occlusion of the connector with the thumb. A choking response, regardless of thumb-occlusion, irregular breaths, or other signs of difficulty breathing are indicative of angiocatheter malpositioning and usually indicate esophageal intubation. CRITICAL STEP Failure to quickly reverse an esophageal intubation can be lethal. If esophageal intubation is suspected, the catheter should be quickly removed and reinserted once the animal has resumed a normal breathing pattern. Etomidate is the anesthetic of choice as, of all available rodent anesthetics, this agent provokes the least cardiovascular toxicity (hypotension, arrhythmia, cardiac arrest).</li>
	<li>Lower plethysmograph table until parallel with the work bench and turn subject 180&deg; until facing air ventilator port. Turn animal on its side before connecting to the ventilator.</li>
	<li>A successful intubation is further confirmed when, after securing an air-tight connection and activating the ventilator (functioning at 150 breaths/minute, 9 ml/g tidal volume, 100% oxygen), thoracoabdominal excursion is seen to pace with the ventilator.</li>
</ol>
Intravenous line:
<ol start="13">
	<li>A 10 mm, 27ga needle is removed from its syringe-connector by way of melting it free, and bending the needle 90&deg; at the midpoint using sterile forceps and hemostat so that the bevel faces into the angle. The non-beveled end is connected to the PE10 tubing leading to the IV injection port.</li>
	<li>To prevent potentially fatal air embolization, the tubing and needle are purged with 37&deg;C, 0.9% NaCl via the 1 ml syringe. The injection port consists of a 27ga needle, pushed through a hole drilled into the cap of a 15ml centrifuge tube. The cap is filled with saline such that the end of the needle is constantly submerged, thus reducing the likelihood that air will be entrained in the needle and injected intravenously.</li>
	<li>With the mouse remaining under the heat lamp, the needle is aligned at the caudal extreme of the tail parallel to and over the lateral vein. The needle is run slightly beneath the skin, while directed cranially along the vein s length and pushed subcutaneously to the bend. Successful IV placement is confirmed by observing blood backflow into the IV tubing with slight pulling of the syringe plunger. Furthermore, there should be unimpeded flow through the IV line upon injection of 50-100 &mu;l saline into the tail vein. Occasional tail veins cannot be stably cannulated. In these instances, the mouse can be rotated 180 degrees to the other side and the other tail IV usually accessed without difficulty.</li>
	<li>After removing the heat lamp from the setup, the subject is enclosed in the plethysmograph, thereafter secured as air-tight with the application of 4 clamps. CRITICAL STEP Allowing the heat lamp to remain on will heat the air in the plethysmograph chamber and potentially alter subsequent measurements of RRS Sterility of i.v. needles and solutions must be maintained. Sterilization of needles is accomplished by immersing and flushing with 70% ethanol followed by rinsing and flushing with sterile saline solution prior to i.v. insertion. Moreover, the tail should be cleaned with 70% ethanol or isopropyl alcohol prior to i.v. insertion.</li>
</ol>
Airway resistance measurements:
<ol start="17">
	<li>Peak resistance is determined by continuous quantitation of the quotient DPt/V (where DPt is the change in the tracheal pressure and V is air flow) at points of equal lung volume (70% tidal volume). DPt is determined by using a pressure transducer connected to the tracheal angiocatheter. To determine V, plethysmograph pressure variations are calibrated to changes in volume over the physiologic ranges studied. The differential of plethysmograph volume over time, as computed by the preamp module, is V. After establishing a stable baseline RRS (&lt;5% variation over 3 minutes), five successive doses (volume = 2 &mu;l/g body weight) of increasing concentrations of acetylcholine chloride (0.058, 0.18, 0.59, 1.58 and 5.8 mg/kg of body weight, in 0.9% saline at pH 7.4, maintained on ice and hand-warmed prior to each injection) are injected over one second via the i.v., with each subsequent dose administered upon return of RRS to baseline, until a tripling of baseline resistance (approximately 12 cm H2O x ml-1 x sec, i.e., a 200% increase in airway resistance above the typical baseline of approximately 4 cm H2O x ml-1 x sec) is achieved. The provocative concentration of Ach, in mg/g of body weight, that causes a 200% increase in RRS from baseline values (termed PC200), is computed by mathematical interpolation of the Ach-RRS dose-response curves.</li>
	<li>Once PC200 values have been reached, release fasteners, and dismantle the plethysmograph. A maximum of 5 increasing doses of Ach is given. The Ach concentration range given above is suitable for achieving PC200 values for most naive mouse strains. CRITICAL STEP When a baseline of approximately 4 cm H2O x ml-1 x sec is established on the resistance monitor for 30 seconds, a 60 &mu;L of normal saline can be injected i.v. to confirm that the appropriate plane of anesthesia has been achieved. With complete anesthesia, there will be no significant change in resistance; an increase in resistance or movement of limbs or tail represents a sign of physical distress and indicates the need for additional anesthetic.</li>
	<li>Remove IV from the tail vein and then disconnect the animal from the ventilator, maintaining a patent airway by keeping the tracheal cannula in place. Occasional animals fail to resume spontaneous breathing immediately. In these instances, respiration can be encouraged by gently massaging the thorax. CRITICAL STEP Spontaneous breathing must be established prior to transfer to the recovery chamber; otherwise deaths will occur.</li>
	<li>Upon resumption of spontaneous breathing, mice are transferred with tracheal cannulae in place, to a chamber purged with 100% O2 and maintaining at 37&deg;C using a heat lamp. Within 15-20 min, mice are breathing strongly and beginning to move their extremities, at which point the tracheal catheter can be removed and animals safely transferred to their regular cages. CRITICAL STEP The airway is readily obstructed in the unconscious mouse due to acetylcholine-induced hyper-salivation and is the major reason for asphyxiation-related death in anesthetized mice following airway physiologic measurements. For this reason, the tracheal cannula must remain in place, even in mice not undergoing bronchoalveolar lavage, until they are arousable and should not be removed until hyper-salivaton has ceased.</li>
</ol>
Bronchoalveolar lavage:
<ol start="21">
	<li>Collection of bronchoalveolar lavage fluid is safe when mice recover sufficiently their gag reflex (~20 min after placement in recovery chamber). The gag reflex is assessed by gently sliding the angiocatheter inward and outward; obvious coughing or struggling indicate that the gag reflex has returned. CRITICAL STEP Allowing too prolonged a recovery time will greatly decrease the efficiency of BAL return from individual mice; thus, the gag reflex must be monitored every few minutes, following the suggested 20 min resting period. If mice are unable to tolerate the lavage procedure due to partial awakening, 3.2% isoflurane vapor anesthesia can be used.</li>
	<li>A metallic intubation guide wire (0.5mm O.D.), with a continuous bend of ~30&deg; directed to the left lobe of the lung, is inserted into the angiocatheter. The guide wire and angiocatheter are advanced together into the left lobe of the lung, such that the catheter (hub excluded) extends beyond the front teeth by only 1mm. CRITICAL STEP Failure to isolate left lung will greatly reduce yield, while enhancing the likelihood of animal death. Care must be taken to ensure that the tip of the guide wire does not pass through the open end of the angiocatheter. Advancing the angiocathether with the metallic tip protruding could lead a tracheal laceration and death due to tracheal rupture.</li>
	<li>Keeping the angiocatheter in place, the guidewire is removed and 300 &mu;L of PBS (pH 7.4, sterile) is flushed into the left lung via a 1 ml syringe. Immediately after, while drawing up the syringe plunger to create negative pressure, the angiocatheter is slowly (3 s) removed while intensely massaging the lung. A BAL return of 100-200 &mu;L is expected.</li>
	<li>Immediately return lavaged mice to the 37&deg;C, 100% O2 chamber while continuously massaging the thorax. Place mice on their left side until fully recovered (~20 min). The animals are then placed back into their cages.</li>
</ol>
TIMING:
Per mouse, the entire procedure should take no longer than 1 hour to accomplish: Step 3-4, 5-10 min.; Steps 5-21, 10 min.; Step 22, 20-30 min.; Steps 23-24, 10 min. With increased proficiency and by staggering subjects in the protocol, up to 3 mice/hour may be processed. 
Representative Results:
Airway hyperreactivity in mice, as determined by measures of PC200 values, is a consequence of activation and recruitment to the lungs of T cells and secretion of the cytokine IL-135-7. Thus, airway hyperreactivity is not the inevitable consequence of airway challenge with allergen, but rather depends on an intact T cell immune compartment and the time required for T cell responses to develop in the setting of repeated allergen exposure. As shown in Fig. 2a, airway hyperreactivity, defined as PC200 values that are significantly lower compared to baseline values, developed after 5 allergen challenges with no further significant increase after the sixth challenge. For reasons that are not fully understood, airway reactivity decreased (PC200 values increased) after the first allergen challenge (Fig. 2a). Similar trends are apparent by comparing Ach dose response curves for the same mice (Fig. 2b). However, it is apparent here that full airway hyperreactivity develops abruptly after the fifth allergen challenge, such that mice become more than 30-fold more sensitive to Ach between the fourth and sixth challenges. Together, these findings indicate that the most reliable measurements of AHR are obtained after six allergen challenges (12 days); measurements at earlier timepoints are likely to yield highly variable data. Mice repeatedly challenged with vehicle intranasally (saline) do not develop airway hyperreactivity, and, at all doses of Ach given, RRS measurements do not significantly vary from baseline values (Fig. 3 and data not shown).
Prior to the emergence of robust AHR, the dominant cell type of the airway induced by allergen was the neutrophil (Fig. 4). Similar to the trend for AHR, however, eosinophilia gradually strengthened with repeated allergen challenge and the eosinophil became the numerically dominant cell type in BAL fluid after the sixth challenge, coincident with a marked decline in neutrophis numbers (Fig. 4). Macrophages initially increased in number with the first few allergen challenges and fluctuated in abundance thereafter. Lymphocyte abundance did not change significantly regardless of the number of allergen challenges and, paradoxically given their primary importance to the model, are typically the least numerous cell in the BAL fluid.
Airway resistance measurements in mice receiving neither allergen challenge nor BAL sampling did not vary over the 17 days of experimentation. Repeated BAL fluid sampling in the absence of airway physiology measurements or allergen challenge were also performed, and showed only an enhanced neutrophil and macrophage recruitment to the airways that did not persist beyond 5 days (data not shown). These findings demonstrate that the prominent neutrophilia observed in allergen challenged mice is largely the result of the procedure and not the antigen.
In control, PBS-challenged mice, airway resistance measurements also did not vary significantly over time. Enhanced macrophage and neutrophil, but not eosinophil, recruitment to BAL fluid was also seen in these mice, similar to those changes observed in mice receiving only repeated BAL fluid sampling (Fig. 4 b, d). Together, these data underscore the importance of the allergen, and not the different manipulations of the airway, to the induction of both allergic (eosinophilic) airway inflammation and AHR. 
Similar results can be expected using intranasal allergens analogous to the proteinase that we have used here. However, many investigators use ovalbumin to induce allergic lung disease. After an appropriate period of intradermal or intraperitoneal priming (1-2 weeks) with ovalbumin precipitated in an aluminum salt, a robust asthma phenotype, including airway hyperreactivity, can be expected within 24 hours after a single intranasal challenge with soluble ovalbumin.
<img src="/files/ftp_upload/1720/1720fig1.jpg" alt="Figure 1" /> Figure 1. Photographic representation of a rodent plethysmograph, immediately prior to airway physiology measurement recording.
<img src="/files/ftp_upload/1720/1720fig2.jpg" alt="Figure 2" /> Figure 2. Airway resistance measurements. A) For statistical purposes, antilog PC200 values are reported. Note the large increase in the antilog PC200 after first challenge and subsequent decrease following further challenges. B) Respiratory system resistance (RRS): Note the steepness of the Ach-RRS dose-response curves after sixth and seventh challenges. Error bars represent SEM.
<img src="/files/ftp_upload/1720/1720fig3.jpg" alt="Figure 3" /> Figure 3. Representative real-time respiratory system resistance (RRS) tracings from a na&iuml;ve (A) and 6X allergen challenged mouse (B) receiving consecutive IV doses of Ach. Dose values are presented in mg/kg units.
<img src="/files/ftp_upload/1720/1720fig4.jpg" alt="Figure 4" /> Figure 4. Differential immune cell counts in the bronchoalveolar lavage samples derived from the left lungs of mice treated with 7 consecutive intranasal challenges. Percent (%) abundance of immune cells in mice treated with allergen (A) or PBS (B). Total number of immune cells from mice treated with allergen (C) or PBS (D). Values represented as mean +/- sem.

<ol>
	<li>Hamelmann, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Noninvasive+measurement+of+airway+responsiveness+in+allergic+mice+using+barometric+plethysmography.">Noninvasive measurement of airway responsiveness in allergic mice using barometric plethysmography.</a> Am. J. Respir. Crit. Care Med. 156, 766-775 (1997).</li><li>Adler, A., Cieslewicz, G., Irvin, C. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Unrestrained+plethysmography+is+an+unreliable+measure+of+airway+responsiveness+in+BALB/c+and+C57BL/6+mice.">Unrestrained plethysmography is an unreliable measure of airway responsiveness in BALB/c and C57BL/6 mice.</a> J. Appl. Physiol. 97, 286-292 (2004).</li><li>Bates, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+use+and+misuse+of+penh+in+animal+models+of+lung+disease.">The use and misuse of penh in animal models of lung disease.</a> Am. J. Respir. Cell Mol. Biol. 31, 373-374 (2004).</li><li>Lundblad, L. K., Irvin, C. G., Adler, A., Bates, J. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+reevaluation+of+the+validity+of+unrestrained+plethysmography+in+mice.">A reevaluation of the validity of unrestrained plethysmography in mice.</a> J. Appl. Physiol. 93, 1198-1207 (2002).</li><li>Grunig, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Requirement+for+IL-13+independently+of+IL-4+in+experimental+asthma.">Requirement for IL-13 independently of IL-4 in experimental asthma.</a> Science. 282, 2261-2263 (1998).</li><li>Corry, D. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Requirements+for+allergen-induced+airway+hyperreactivity+in+T+and+B+cell-deficient+mice.">Requirements for allergen-induced airway hyperreactivity in T and B cell-deficient mice.</a> Mol. Med. 4, 344-355 (1998).</li><li>Corry, D. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Interleukin+4,+but+not+interleukin+5+or+eosinophils,+is+required+in+a+murine+model+of+acute+airway+hyperreactivity.">Interleukin 4, but not interleukin 5 or eosinophils, is required in a murine model of acute airway hyperreactivity.</a> J. Exp. Med. 183, 109-117 (1996).</li><li>Amdur, M. O., Mead, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mechanics+of+respiration+in+unanesthetized+guinea+pigs.">Mechanics of respiration in unanesthetized guinea pigs.</a> Am J Physiol. 192, 364-368 (1958).</li></ol>

The study of asthma, and various other airway obstructive diseases, constitutes an active and expanding field of biomedical research. An important component of asthma-related experimental research is the capacity to measure changes in airway size under varying conditions. Excessive airway narrowing in response to provocative challenge, a canonical feature of asthma and related lung diseases and a property of the airway termed airway hyperresponsiveness, is a major component of clinically significant attacks leading to sh...

Airway physiology measurement software (Rescomp) was custom prepared (Millenium Premier Group; 415-519-4371). Data was analyzed using a PC workstation running Windows XP equipped with a Pentium III CPU (Intel, Inc. Santa Clara, CA) and a 17-pin analog to digital signal converter (National Instruments, #PC-LPM16). A small animal airway physiology workstation was custom assembled (Millenium Premier Group) using commercially available pressure transducers (part #TRD5700 and TRD4510), preamp modules (part #MAX2270), chassis (part # MAX1320; all from Buxco, Inc. Wilmington, NC) and a customized small animal plethysmograph. 0.5mm external diameter fiber-optic thread, connected to light source (Cole Palmer Illuminator, 41722 series) Ventilator (Harvard Apparatus Mouse Ventilator, #687) 10 mm, 27ga needle (BD Biosciences, cat. no. 309602) Heat lamp 1 ml syringe (BD Biosciences, cat. no. 305109) 4 clamps (Pony 3200 spring clamp) 0.5 mm external wire for intubation guide Hemacytometer Superfrost/plus microscope slides (Fisher cat. no. 12-550-15) Shandon Filter Cards (Thermo cat. no. 5991022) Differential cell slide stain (Fisher cat. no. 122911) Light microscope (Leica) Cytospin 3 (Shandon) 20 ga, 1.25 inch ProtectIV intravenous catheters (Smith Medical) 0.5 mm polymer optical fiber (Edmund Optics # NT02-532).

a reversible, non-invasive method for airway resistance measurements and bronchoalveolar lavage fluid sampling in mice

Airway hyperreactivity (AHR) measurements and bronchoalveolar lavage (BAL) fluid sampling are essential to experimental asthma models, but repeated procedures to obtain such measurements in the same animal are generally not feasible.  Here, we demonstrate protocols for obtaining from mice repeated measurements of AHR and bronchoalveolar lavage fluid samples.  Mice were challenged intranasally seven times over 14 days with a potent allergen or sham treated.  Prior to the initial challenge, and within 24 hours following each intranasal challenge, the same animals were anesthetized, orally intubated and mechanically ventilated.  AHR, assessed by comparing dose response curves of respiratory system resistance (RRS) induced by increasing intravenous doses of acetylcholine (Ach) chloride between sham and allergen-challenged animals, were determined.  Afterwards, and via the same intubation, the left lung was lavaged so that differential enumeration of airway cells could be performed.  These studies reveal that repeated measurements of AHR and BAL fluid collection are possible from the same animals and that maximal airway hyperresponsiveness and airway eosinophilia are achieved within 7-10 days of initiating allergen challenge.  This novel technique significantly reduces the number of mice required for longitudinal experimentation and is applicable to diverse rodent species, disease models and airway physiology instruments.  

Watch this Scientific Journal Video about A Reversible, Non-invasive Method for Airway Resistance Measurements and Bronchoalveolar Lavage Fluid Sampling in Mice at JoVE.com

A Reversible, Non-invasive Method for Airway Resistance Measurements and Bronchoalveolar Lavage Fluid Sampling in Mice

Repeated measurements of rodent respiratory physiology and sampling of airway inflammatory cells are desirable, but generally not feasible. Here we describe a repeatable method for orally intubating mice that permits repeated measurements of airway hyperreactivity and sampling of airway inflammatory cells.

Airway hyperreactivity (AHR) measurements and bronchoalveolar lavage (BAL) fluid sampling are essential to experimental asthma models, but repeated ...

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22.3K Views.  Baylor College of Medicine (BCM). Repeated measurements of rodent respiratory physiology and sampling of airway inflammatory cells are desirable, but generally not feasible. Here we describe a repeatable method for orally intubating mice that permits repeated measurements of airway hyperreactivity and sampling of airway inflammatory cells.

Video: A Reversible, Non-invasive Method for Airway Resistance Measurements and Bronchoalveolar Lavage Fluid Sampling in Mice

Pyrosequencing: A Simple Method for Accurate Genotyping

Pharmacogenetic research benefits first-hand from the abundance of information provided by the completion of the Human Genome Project. With such a tremendous amount of data available comes an explosion of genotyping methods. Pyrosequencing(R) is one of the most thorough yet simple methods to date used to analyze polymorphisms. It also has the ability to identify tri-allelic, indels, short-repeat polymorphisms, along with determining allele percentages for methylation or pooled sample assessment. In addition, there is a standardized control sequence that provides internal quality control. This method has led to rapid and efficient single-nucleotide polymorphism evaluation including many clinically relevant polymorphisms. The technique and methodology of Pyrosequencing is explained.


Pyrosequencing(R) is one of the most thorough yet simple methods to date used to analyze polymorphisms. This method has led to rapid and efficient single-nucleotide polymorphism evaluation including many clinically relevant polymorphisms. The technique and methodology of Pyrosequencing is explained.

Pharmacogenetic research benefits first-hand from the abundance of information provided by the completion of the Human Genome Project. With such a ...

Labeling hESCs and hMSCs with Iron Oxide Nanoparticles for Non-Invasive in vivo Tracking with MR Imaging

In recent years, stem cell research has led to a better understanding of developmental biology, various diseases and its potential impact on regenerative medicine. A non-invasive method to monitor the transplanted stem cells repeatedly in vivo would greatly enhance our ability to understand the mechanisms that control stem cell death and identify trophic factors and signaling pathways that improve stem cell engraftment. MR imaging has been proven to be an effective tool for the in vivo depiction of stem cells with near microscopic anatomical resolution. In order to detect stem cells with MR, the cells have to be labeled with cell specific MR contrast agents. For this purpose, iron oxide nanoparticles, such as superparamagnetic iron oxide particles (SPIO), are applied, because of their high sensitivity for cell detection and their excellent biocompatibility. SPIO particles are composed of an iron oxide core and a dextran, carboxydextran or starch coat, and function by creating local field inhomogeneities, that cause a decreased signal on T2-weighted MR images. This presentation will demonstrate techniques for labeling of stem cells with clinically applicable MR contrast agents for subsequent non-invasive in vivo tracking of the labeled cells with MR imaging.

For the evaluation of new stem cell therapies it is important to non-invasively track the injected cells in vivo. This video will show you how to label human mesenchymal and embryonic stem cells with iron oxide based contrast agents in vivo for subsequent MR imaging in vivo.

In recent years, stem cell research has led to a better understanding of developmental biology, various diseases and its potential impact on regenerative ...

Labeling Stem Cells with Fluorescent Dyes for non-invasive Detection with Optical Imaging

Optical imaging (OI) is an easy, fast and inexpensive tool for in vivo monitoring of new stem cell based therapies. The technique is based on ex vivo labeling of stem cells with a fluorescent dye, subsequent intravenous injection of the labeled cells and visualization of their accumulation in specific target organs or pathologies. The presented technique demonstrates how we label human mesenchymal stem cells (hMSC) by simple incubation with the lipophilic fluorescent dye DiD (C67H103CIN2O3S) and how we label human embryonic stem cells (hESC) with the FDA approved fluorescent dye Indocyanine Green (ICG). The uptake mechanism is via adherence and diffusion of the lypophilic dye across the phospholipid cell membrane bilayer. The labeling efficiency is usually improved if the cells are incubated with the dye in serum-free media as opposed to incubation in serum-containing media. Furthermore, the addition of the transfection agent Protamine Sulfate significantly improves contrast agent uptake.

This video shows techniques for labeling of human embryonic stem cells and mesenchymal stem cells with fluorescent dyes. This technique can be used for an in vivo tracking of transplanted stem cells with optical imaging and for histopathological correlations with fluorescence microscopy.

Optical imaging (OI) is an easy, fast and inexpensive tool for in vivo monitoring of new stem cell based therapies. The technique is based on ex vivo ...

Seawater Sampling and Collection

This video documents methods for collecting coastal marine water samples and processing them for various downstream applications including biomass concentration, nucleic acid purification, cell abundance, nutrient and trace gas analyses. For today's demonstration samples were collected from the deck of the HMS John Strickland operating in Saanich Inlet. An A-frame derrick, with a multi-purpose winch and cable system, is used in combination with Niskin or Go-Flo water sampling bottles. Conductivity, Temperature, and Depth (CTD) sensors are also used to sample the underlying water mass. To minimize outgassing, trace gas samples are collected first.  Then, nutrients, water chemistry, and cell counts are determined. Finally, waters are collected for biomass filtration. The set-up and collection time for a single cast is ~1.5 hours at a maximum depth of 215 meters. Therefore, a total of 6 hours is generally needed to complete the collection series described here.

This video documents methods for collecting coastal marine water samples and processing them for various downstream applications including biomass concentration, nucleic acid purification, cell abundance, nutrient and trace gas analyses.

This video documents methods for collecting coastal marine water samples and processing them for various downstream applications including biomass ...

A 3D System for Culturing Human Articular Chondrocytes in Synovial Fluid

Cartilage destruction is a central pathological feature of osteoarthritis, a leading cause of disability in the US. Cartilage in the adult does not regenerate very efficiently in vivo; and as a result, osteoarthritis leads to irreversible cartilage loss and is accompanied by chronic pain and immobility 1,2. Cartilage tissue engineering offers promising potential to regenerate and restore tissue function. This technology typically involves seeding chondrocytes into natural or synthetic scaffolds and culturing the resulting 3D construct in a balanced medium over a period of time with a goal of engineering a biochemically and biomechanically mature tissue that can be transplanted into a defect site in vivo 3-6. Achieving an optimal condition for chondrocyte growth and matrix deposition is essential for the success of cartilage tissue engineering. 
 In the native joint cavity, cartilage at the articular surface of the bone is bathed in synovial fluid. This clear and viscous fluid provides nutrients to the avascular articular cartilage and contains growth factors, cytokines and enzymes that are important for chondrocyte metabolism 7,8. Furthermore, synovial fluid facilitates low-friction movement between cartilaginous surfaces mainly through secreting two key components, hyaluronan and lubricin 9 10. In contrast, tissue engineered cartilage is most often cultured in artificial media. While these media are likely able to provide more defined conditions for studying chondrocyte metabolism, synovial fluid most accurately reflects the natural environment of which articular chondrocytes reside in.
 Indeed, synovial fluid has the advantage of being easy to obtain and store, and can often be regularly replenished by the body. Several groups have supplemented the culture medium with synovial fluid in growing human, bovine, rabbit and dog chondrocytes, but mostly used only low levels of synovial fluid (below 20&#37;) 11-25. While chicken, horse and human chondrocytes have been cultured in the medium with higher percentage of synovial fluid, these culture systems were two-dimensional 26-28. Here we present our method of culturing human articular chondrocytes in a 3D system with a high percentage of synovial fluid (up to 100&#37;) over a period of 21 days. In doing so, we overcame a major hurdle presented by the high viscosity of the synovial fluid. This system provides the possibility of studying human chondrocytes in synovial fluid in a 3D setting, which can be further combined with two other important factors (oxygen tension and mechanical loading) 29,30 that constitute the natural environment for cartilage to mimic the natural milieu for cartilage growth. Furthermore, This system may also be used for assaying synovial fluid activity on chondrocytes and provide a platform for developing cartilage regeneration technologies and therapeutic options for arthritis.

A 3D system of culturing human articular chondrocytes in high levels of synovial fluid is described. Synovial fluid reflects the most natural microenvironment for articular cartilage, and can be easily obtained and stored. This system thus can be used for studying cartilage regeneration and for screening therapeutics for treating arthritis.

Cartilage destruction is a central pathological feature of osteoarthritis, a leading cause of disability in the US. Cartilage in the adult does not ...

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 ...

Ex vivo Method for High Resolution Imaging of Cilia Motility in Rodent Airway Epithelia

An ex vivo technique for imaging mouse airway epithelia for quantitative analysis of motile cilia function important for insight into mucociliary clearance function has been established. Freshly harvested mouse trachea is cut longitudinally through the trachealis muscle and mounted in a shallow walled chamber on a glass-bottomed dish. The trachea sample is positioned along its long axis to take advantage of the trachealis muscle to curl longitudinally. This allows imaging of ciliary motion in the profile view along the entire tracheal length. Videos at 200 frames/sec are obtained using differential interference contrast microscopy and a high speed digital camera to allow quantitative analysis of cilia beat frequency and ciliary waveform. With the addition of fluorescent beads during imaging, cilia generated fluid flow also can be determined. The protocol time spans approximately 30 min, with 5 min for chamber preparation, 5-10 min for sample mounting, and 10-15 min for videomicroscopy.

Ex vivo Method for High Resolution Imaging of Cilia Motility in Rodent Airway Epithelia

An easy and reliable technique for visualizing and quantifying airway cilia motility and cilia generated flow using mouse trachea is described. This technique can be modified to determine how a wide range of factors influence cilia motility, including pharmacological agents, genetic factors, environmental exposures, and/or mechanical factors such as mucus load.

An ex vivo technique for imaging mouse airway epithelia for quantitative  analysis of motile cilia function important for insight into mucociliary  ...

Functional Reconstitution and Channel Activity Measurements of Purified Wildtype and Mutant CFTR Protein

The Cystic Fibrosis Transmembrane Conductance Regulator (CFTR) is a unique channel-forming member of the ATP Binding Cassette (ABC) superfamily of transporters. The phosphorylation and nucleotide dependent chloride channel activity of CFTR has been frequently studied in whole cell systems and as single channels in excised membrane patches. Many Cystic Fibrosis-causing mutations have been shown to alter this activity. While a small number of purification protocols have been published, a fast reconstitution method that retains channel activity and a suitable method for studying population channel activity in a purified system have been lacking. Here rapid methods are described for purification and functional reconstitution of the full-length CFTR protein into proteoliposomes of defined lipid composition that retains activity as a regulated halide channel. This reconstitution method together with a novel flux-based assay of channel activity is a suitable system for studying the population channel properties of wild type CFTR and the disease-causing mutants F508del- and G551D-CFTR. Specifically, the method has utility in studying the direct effects of phosphorylation, nucleotides and small molecules such as potentiators and inhibitors on CFTR channel activity. The methods are also amenable to the study of other membrane channels/transporters for anionic substrates.

Described here is a rapid and effective procedure for functional reconstitution of purified wild-type and mutant CFTR protein that preserves activity for this chloride channel, which is defective in Cystic Fibrosis. Iodide efflux from reconstituted proteoliposomes mediated by CFTR allows studies of channel activity and the effects of small molecules.

The Cystic Fibrosis Transmembrane Conductance Regulator (CFTR) is a unique channel-forming member of the ATP Binding Cassette (ABC) superfamily of ...

Fecal Glucocorticoid Analysis: Non-invasive Adrenal Monitoring in Equids

Adrenal activity can be assessed in the equine species by analysis of feces for corticosterone metabolites. During a potentially aversive situation, corticotrophin releasing hormone (CRH) is released from the hypothalamus in the brain. This stimulates the release of adrenocorticotrophic hormone (ACTH) from the pituitary gland, which in turn stimulates release of glucocorticoids from the adrenal gland. In horses the glucocorticoid corticosterone is responsible for several adaptations needed to support equine flight behaviour and subsequent removal from the aversive situation. Corticosterone metabolites can be detected in the feces of horses and assessment offers a non-invasive option to evaluate long term patterns of adrenal activity. Fecal assessment offers advantages over other techniques that monitor adrenal activity including blood plasma and saliva analysis. The non-invasive nature of the method avoids sampling stress which can confound results. It also allows the opportunity for repeated sampling over time and is ideal for studies in free ranging horses. This protocol describes the enzyme linked immunoassay (EIA) used to assess feces for corticosterone, in addition to the associated biochemical validation.

Adrenal activity can be assessed in the equine species by analysis of feces for corticosterone metabolites. The method offers a non-invasive option to assess long term patterns in both domestic and free ranging horses. This protocol describes the enzyme linked immunoassay involved and the associated biochemical validation.

Adrenal activity can be assessed in the equine species by analysis of feces for corticosterone metabolites. During a potentially aversive situation, ...

The Assembly and Application of 'Shear Rings': A Novel Endothelial Model for Orbital, Unidirectional and Periodic Fluid Flow and Shear Stress

Deviations from normal levels and patterns of vascular fluid shear play important roles in vascular physiology and pathophysiology by inducing adaptive as well as pathological changes in endothelial phenotype and gene expression. In particular, maladaptive effects of periodic, unidirectional flow induced shear stress can trigger a variety of effects on several vascular cell types, particularly endothelial cells. While by now endothelial cells from diverse anatomic origins have been cultured, in-depth analyses of their responses to fluid shear have been hampered by the relative complexity of shear models (e.g., parallel plate flow chamber, cone and plate flow model). While these all represent excellent approaches, such models are technically complicated and suffer from drawbacks including relatively lengthy and complex setup time, low surface areas, requirements for pumps and pressurization often requiring sealants and gaskets, creating challenges to both maintenance of sterility and an inability to run multiple experiments. However, if higher throughput models of flow and shear were available, greater progress on vascular endothelial shear responses, particularly periodic shear research at the molecular level, might be more rapidly advanced. Here, we describe the construction and use of shear rings: a novel, simple-to-assemble, and inexpensive tissue culture model with a relatively large surface area that easily allows for a high number of experimental replicates in unidirectional, periodic shear stress studies on endothelial cells.

The Assembly and Application of &#039;Shear Rings&#039;: A Novel Endothelial Model for Orbital, Unidirectional and Periodic Fluid Flow and Shear Stress

Different levels and patterns of fluid shear are known to modulate endothelial gene expression, phenotype and susceptibility to disease. We discuss the assembly and use of 'shear rings': a model that produces unidirectional, periodic shear stress patterns. Shear rings are simple to assemble, economical and can produce high cell yields.

Deviations from normal levels and patterns of vascular fluid shear play important roles in vascular physiology and pathophysiology by inducing adaptive as ...