This work was funded, in part, by NIH grants HL087653 and HL105314. Some of the methods were developed in the laboratory of Dr. Robert Mecham at the Washington University School of Medicine.

1. Ultrasound and pressure system
<ol>
	<li>Start the ultrasound system according to the manufacturers instructions (Vevo 770, Visualsonics). Fill the appropriate probe (age E18 - P7 = model 708, age &gt; P7 = model 707B) with distilled water and connect to the ultrasound system. Place the probe in the adjustable stand at the approximate orientation necessary to obtain an LV long-axis view for a mouse mounted on the imaging platform so that the LV apex is pointed toward the injection arm.</li>
	<li>The pressure system consists of a fluid-filled pressure transducer, bridge amplifier, data acquisition system and data recording software (LabChart). The connections consist of three-way stopcocks, tubing, luer locks and hose barbs. A 20 mL syringe is filled with 10% heparinized saline for flushing at one end. The other end connects to a needle and a 3 mL syringe body is modified for use as a casing to hold the needle tubing in the injection arm (Figure 1).</li>
	<li>Calibrate the pressure transducer system with a manometer.</li>
	<li>Mount the needle tubing and casing in the injection arm of the ultrasound system. Clamp the pressure transducer in a ring stand at the approximate height of the imaging platform. Attach a needle appropriate for the age of the mouse (age E18 - P3 = 30G, age &gt; P3 = 25G) to the needle tubing. Smaller needles get clogged easier, are more difficult to advance through the chest wall, and have a slower response time. Hence, the largest needle size that fits in the LV lumen is used for each age.</li>
	<li>Place a mound of ultrasound gel on the imaging platform and carefully line up the probe using the adjustable stand so that the needle can be advanced directly into the gel under the center of the imaging probe (Figure 2). The needle must be far enough below the probe to not puncture the probe cover, but close enough to be within the probe focal depth. If necessary, the needle may be bent by hand so that it remains in the correct plane throughout its travel.</li>
	<li>Maintaining alignment, retract the needle and raise the imaging probe to place a mouse on the imaging platform. Flush the needle to eliminate any gel that may have entered the tip.</li>
</ol>
2. Mouse preparation
<ol>
	<li>All methods shown in this protocol have been approved by the Institutional Animal Care and Use Committee. Anesthetize a timed pregnant mother (E18) or neonatal mouse (P1 - 20) by continuous inhalation of 1.5% isoflurane. Tubing can be inserted into a standard nose cone to accommodate the small head of an early neonatal mouse (P1 - 15).</li>
	<li>Secure the mouse to the imaging platform of the ultrasound system in a dorsal decubitus position with the LV apex pointing toward the injection arm. For pregnant mothers, apply ultrasound gel to monitor heart rate with the attached ECG detector and insert the rectal thermometer for feedback control of the body temperature. Neonatal mice are too small for these detectors, so heart rate is monitored during the pressure measurements and body temperature is maintained with an external heat lamp.</li>
	<li>Remove hair from the abdomen of the pregnant mother or chest of the young mouse with a depilatory cream. Neonatal mice (P1 - 10) do not require any hair removal.</li>
	<li>For pregnant mothers, cut open the abdomen using dissecting scissors and curved forceps. Externalize one uterine horn and place on gauze moistened with warm saline solution. Carefully manipulate an embryo into the proper position to obtain a parasternal long axis view of the LV using the ultrasound probe. Measure the pressure in all embryos on this side before externalizing the other uterine horn. Keep all embryos and the abdomen of the mother moist with periodic addition of warm saline.</li>
</ol>
3. Pressure measurement
<ol>
	<li>Place prewarmed, degassed ultrasound gel on the embryonic or neonatal mouse. Neonatal mice that have not been depilated should have a small amount of ultrasound gel rubbed onto their chest before the remaining ultrasound gel is applied to enhance coupling. Lower the imaging probe and adjust the angle of the imaging platform to obtain an optimal parasternal LV long axis view. Do not adjust the angle of the probe because the needle alignment will be lost.</li>
	<li>While monitoring the LV image on the ultrasound screen, slowly advance the needle until it is near the bottom of the probe. Adjust the needle and probe position, if necessary, to correct alignment shifts. Gently flush the needle with saline. Start recording pressure data and zero the pressure transducer with the LabChart software. The transducer drifts with temperature and position, so it is important to zero it just before measuring pressure in each mouse.</li>
	<li>Slowly advance the needle until it can be seen at the edge of the ultrasound image, but not yet in the LV. Adjust the needle position, if necessary, to enter the LV at the apex. Adjust the probe position, if necessary, to maintain a clear image of the needle. Advance the needle into the LV lumen (Figure 3), while recording pressure data and monitoring the needle location on the ultrasound image. The start of pulsatile recordings from the pressure transducer confirms the location inside the LV lumen (Figure 4).</li>
	<li>If the needle appears to be in the LV lumen, but pressure recordings are not pulsatile, flush with a light tap on the syringe plunger, just until a pressure increase is detected (about 5 - 10 &mu;L volume). This should clear any clogs that may be preventing an accurate reading. If pressure readings are still not pulsatile, the needle may be above or below the correct plane and not be inserted into the LV lumen. Retract the needle, flush with saline, and advance again. If pulsatile readings are still not obtained, move on to the next mouse. Use a new needle for each mouse.</li>
	<li>Successful pressure readings can be obtained from 75% of the embryonic and neonatal mice attempted. Readings should take 5 - 10 minutes each and the pressure data for a mouse should not be included if the heart rates are significantly different than expected (Figure 5A). Heart rate has a significant effect on cardiovascular measures and altered heart rates are indicative of cardiac distress. Embryonic distress usually occurs 1 - 1.5 hours after first externalization of the uterus, so speed is important for measuring pressures in an entire litter.</li>
</ol>
4. Representative Results
All results shown are for C57BL6J mice. An image of the needle in the LV lumen for a P1 mouse in shown in Figure 3. The needle is necessary to puncture the chest wall and gain access to the small LV lumen, but it significantly increases the response time of the pressure system. When an approximate step increase in pressure is manually applied to the system, the time to reach 67% of the maximum pressure is .067 sec with the tubing only (most likely indicative of the real time to apply the pressure step), 0.105 sec with a 25 G needle and 0.529 sec with a 30 G needle. The delay in reaching maximum pressure can be seen in the complete tracings shown in Figure 4A and 4C. Although the response time is slower with the 30G needle, the waveform is better captured at the embryonic and early neonatal stages, because heart rate increases with age in mice5. Despite this limitation, the systolic pressure can be calculated assuming that the true LV diastolic (minimum) pressure is zero, and the systolic (maximum) LV pressure is two times the mean LV pressure determined from the readings at steady state (Figure 4B and 4D)6. It is generally assumed that the systolic LV and arterial pressures are equal. The measured heart rates and calculated LV pressures for various ages between E18 and P14 are shown in Figure 5.
<img src="/files/ftp_upload/3756/3756fig1.jpg" /> 
Figure 1. Image of the pressure transducer with attached three-way stopcocks, male (M) and female (F) luer lock connections, hose barbs, tubing, syringes and needle.
<img src="/files/ftp_upload/3756/3756fig2.jpg" /> 
Figure 2. Image of the set up to align the needle with the imaging probe (A). The imaging platform is rotated so that the LV apex of the mouse is facing the injection arm. The probe is mounted in the adjustable stand at the approximate orientation necessary to obtain an LV long-axis image. The needle tubing casing is mounted in the injection arm. The needle is advanced into a mound of ultrasound gel on the imaging platform to determine the proper angle and vertical and horizontal position with respect to the imaging probe (B).
<img src="/files/ftp_upload/3756/3756fig3.jpg" /> 
Figure 3. Image of the needle (N) advanced through the chest wall (CW) and into the LV lumen in a P1 mouse. Scale bar = 0.1 mm.
<img src="/files/ftp_upload/3756/3756fig4.jpg" /> 
Figure 4. Example pulsatile pressure readings as the needle enters the LV of an E18 (A) and P14 (C) mouse. There is a delay in reaching a steady state due to the response time of the system with the needle attached. Zoom views of the readings at steady state are shown in B and D. Note that the minimum E18 LV pressures are near zero, but the full waveform from zero to the maximum pressure cannot be recorded at the higher heart rates of P14 mice. The mean pressures are measured from the steady state readings and it is assumed that LV systolic pressure = 2 x measured mean pressure.
<img src="/files/ftp_upload/3756/3756fig5.jpg" /> 
Figure 5. Measured heart rates (A) and calculated systolic LV pressures (B) for E18 - P14 mice. N = 7 for E18, 5 for P1, 22 for P3, 23 for P7 and 16 for P145-7.

<ol>
	<li>Ji, R. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Onset+of+cardiac+function+during+early+mouse+embryogenesis+coincides+with+entry+of+primitive+erythroblasts+into+the+embryo+proper.">Onset of cardiac function during early mouse embryogenesis coincides with entry of primitive erythroblasts into the embryo proper.</a> Circ. Res. 92 (2), 133-135 (2003).</li><li>Ishiwata, T., Nakazawa, M., Pu, W. T., Tevosian, S. G., Izumo, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Developmental+changes+in+ventricular+diastolic+function+correlate+with+changes+in+ventricular+myoarchitecture+in+normal+mouse+embryos.">Developmental changes in ventricular diastolic function correlate with changes in ventricular myoarchitecture in normal mouse embryos.</a> Circ. Res. 93 (9), 857-865 (2003).</li><li>Huang, Y., Guo, X., Kassab, G. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Axial+nonuniformity+of+geometric+and+mechanical+properties+of+mouse+aorta+is+increased+during+postnatal+growth.">Axial nonuniformity of geometric and mechanical properties of mouse aorta is increased during postnatal growth.</a> Am. J. Physiol. Heart Circ. Physiol. 290 (2), H657-H664 (2006).</li><li>Ishii, T., Kuwaki, T., Masuda, Y., Fukuda, Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Postnatal+development+of+blood+pressure+and+baroreflex+in+mice.">Postnatal development of blood pressure and baroreflex in mice.</a> Auton. Neurosci. 94 (1-2), 34-41 (2001).</li><li>Le, V. P., Knutsen, R. H., Mecham, R. P., Wagenseil, J. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Decreased+aortic+diameter+and+compliance+precedes+blood+pressure+increases+in+postnatal+development+of+elastin-insufficient+mice.">Decreased aortic diameter and compliance precedes blood pressure increases in postnatal development of elastin-insufficient mice.</a> Am. J. Physiol. Heart Circ. Physiol. , (2011).</li><li>Wagenseil, J. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Reduced+vessel+elasticity+alters+cardiovascular+structure+and+function+in+newborn+mice.">Reduced vessel elasticity alters cardiovascular structure and function in newborn mice.</a> Circ. Res. 104 (10), 1217-1224 (2009).</li><li>Wagenseil, J. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+importance+of+elastin+to+aortic+development+in+mice.">The importance of elastin to aortic development in mice.</a> Am. J. Physiol. Heart Circ. Physiol. 299 (2), H257-H264 (2010).</li><li>Ramirez, F., Sakai, L. Y., Rifkin, D. B., Dietz, H. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Extracellular+microfibrils+in+development+and+disease.">Extracellular microfibrils in development and disease.</a> Cell. Mol. Life Sci. 64 (18), 2437-2446 (2007).</li><li>Li, D. Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Elastin+point+mutations+cause+an+obstructive+vascular+disease,+supravalvular+aortic+stenosis.">Elastin point mutations cause an obstructive vascular disease, supravalvular aortic stenosis.</a> Hum. Mol. Genet. 6 (7), 1021-1028 (1997).</li><li>Roth, D. M., Swaney, J. S., Dalton, N. D., Gilpin, E. A., Ross, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Impact+of+anesthesia+on+cardiac+function+during+echocardiography+in+mice.">Impact of anesthesia on cardiac function during echocardiography in mice.</a> Am. J. Physiol. Heart Circ. Physiol. 282 (6), H2134-H2140 (2002).</li><li>Phoon, C. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Imaging+tools+for+the+developmental+biologist:+ultrasound+biomicroscopy+of+mouse+embryonic+development.">Imaging tools for the developmental biologist: ultrasound biomicroscopy of mouse embryonic development.</a> Pediatr. Res. 60 (1), 14-21 (2006).</li></ol>

No conflicts of interest declared.

The protocol presented here provides a method for measuring LV pressure in late embryonic and early neonatal mice. The main limitation of this protocol is the temporal resolution of the pressure system. The pressure signal is damped as it travels from the LV through the needle to the transducer, and only mean pressure values can be recorded. The damping can be minimized by using the largest needle possible, but the needle must fit within the LV lumen for different aged mice. Because the diastolic pressure can be approxim...

<table border="1">
<tbody>
 <tr>
 <td>Name of the reagent/equipment</td>
 <td>Company</td>
 <td>Catalogue number</td>
 <td>Comments</td>
 </tr>
 <tr>
 <td>High resolution ultrasound system</td>
 <td>Visualsonics</td>
 <td>Vevo 770</td>
 <td>Or other appropriate ultrasound 			system</td>
 </tr>
 <tr>
 <td>High frequency ultrasound probes </td>
 <td>Visualsonics</td>
 <td>708 and 707B</td>
 <td>&nbsp;
 </td>
 </tr>
 <tr>
 <td>Imaging platform and injection arm</td>
 <td>Visualsonics</td>
 <td>Imaging Station 2</td>
 <td>With ECG and temperature feedback 			control of platform</td>
 </tr>
 <tr>
 <td>Pressure transducer</td>
 <td>AD Instruments</td>
 <td>MLT 844</td>
 <td>&nbsp;
 </td>
 </tr>
 <tr>
 <td>Bridge amplifier</td>
 <td>AD Instruments</td>
 <td>ML221</td>
 <td>&nbsp;
 </td>
 </tr>
 <tr>
 <td>Data acquisition system</td>
 <td>AD Instruments</td>
 <td>ML866</td>
 <td>&nbsp;
 </td>
 </tr>
 <tr>
 <td>Data recording software</td>
 <td>AD Instruments</td>
 <td>LabChart</td>
 <td>&nbsp;
 </td>
 </tr>
 <tr>
 <td>Ring stand and clamp</td>
 <td>Various suppliers</td>
 <td>&nbsp;
 </td>
 <td>To hold pressure transducer during 			measurements</td>
 </tr>
 <tr>
 <td>3-way stopcocks with luer 			connections, male lock</td>
 <td>Cole Parmer</td>
 <td>30600-02</td>
 <td>&nbsp;
 </td>
 </tr>
 <tr>
 <td>1/16" ID Tygon Tubing</td>
 <td>Cole Parmer</td>
 <td>06408</td>
 <td>&nbsp;
 </td>
 </tr>
 <tr>
 <td>Male and female luers w/ 1/16" hose 			barb</td>
 <td>Cole Parmer</td>
 <td>45510-50 45510-00</td>
 <td>&nbsp;
 </td>
 </tr>
 <tr>
 <td>24" tubing with male and female 			luer at each end</td>
 <td>Cole Parmer</td>
 <td>30600-60</td>
 <td>&nbsp;
 </td>
 </tr>
 <tr>
 <td>3 and 10 mL syringes</td>
 <td>BD Biosciences</td>
 <td>&nbsp;
 </td>
 <td>&nbsp;
 </td>
 </tr>
 <tr>
 <td>30 and 25G needles</td>
 <td>BD Biosciences</td>
 <td>&nbsp;
 </td>
 <td>1.5 inches in length</td>
 </tr>
 <tr>
 <td>Big Ben manometer</td>
 <td>Riester</td>
 <td>1456-100</td>
 <td>&nbsp;
 </td>
 </tr>
 <tr>
 <td>Saline</td>
 <td>Various suppliers</td>
 <td>&nbsp;
 </td>
 <td>&nbsp;
 </td>
 </tr>
 <tr>
 <td>Heparin</td>
 <td>Various suppliers</td>
 <td>&nbsp;
 </td>
 <td>&nbsp;
 </td>
 </tr>
 <tr>
 <td>Ultrasound gel</td>
 <td>Parker</td>
 <td>Aquasonic 100</td>
 <td>&nbsp;
 </td>
 </tr>
 <tr>
 <td>Hair remover lotion</td>
 <td>Nair</td>
 <td>&nbsp;
 </td>
 <td>&nbsp;
 </td>
 </tr>
 </tbody>
</table>

measuring left ventricular pressure in late embryonic and neonatal mice

Blood pressure increases significantly during embryonic and postnatal development in vertebrate animals. In the mouse, blood flow is first detectable around embryonic day (E) 8.51. Systolic left ventricular (LV) pressure is 2 mmHg at E9.5 and 11 mmHg at E14.52. At these mid-embryonic stages, the LV is clearly visible through the chest wall for invasive pressure measurements because the ribs and skin are not fully developed. Between E14.5 and birth (approximately E21) imaging methods must be used to view the LV. After birth, mean arterial pressure increases from 30 - 70 mmHg from postnatal day (P) 2 - 353. Beyond P20, arterial pressure can be measured with solid-state catheters (i.e. Millar or Scisense). Before P20, these catheters are too big for developing mouse arteries and arterial pressure must be measured with custom pulled plastic catheters attached to fluid-filled pressure transducers3 or glass micropipettes attached to servo null pressure transducers4.
Our recent work has shown that the greatest increase in blood pressure occurs during the late embryonic to early postnatal period in mice5-7. This large increase in blood pressure may influence smooth muscle cell (SMC) phenotype in developing arteries and trigger important mechanotransduction events. In human disease, where the mechanical properties of developing arteries are compromised by defects in extracellular matrix proteins (i.e. Marfan's Syndrome8 and Supravalvular Aortic Stenosis9) the rapid changes in blood pressure during this period may contribute to disease phenotype and severity through alterations in mechanotransduction signals. Therefore, it is important to be able to measure blood pressure changes during late embryonic and neonatal periods in mouse models of human disease.
We describe a method for measuring LV pressure in late embryonic (E18) and early postnatal (P1 - 20) mice. A needle attached to a fluid-filled pressure transducer is inserted into the LV under ultrasound guidance. Care is taken to maintain normal cardiac function during the experimental protocol, especially for the embryonic mice. Representative data are presented and limitations of the protocol are discussed.

Watch this Scientific Journal Video about Measuring Left Ventricular Pressure in Late Embryonic and Neonatal Mice at JoVE.com

Measuring Left Ventricular Pressure in Late Embryonic and Neonatal Mice

Measuring left ventricular pressure (LV) in embryonic and neonatal mice is described. Pressure is measured by inserting a needle connected to a fluid-filled transducer into the LV under ultrasound guidance. Care must be taken to maintain normal cardiac function during the experimental protocol.

Blood pressure increases significantly during embryonic and postnatal development in vertebrate animals. In the mouse, blood flow is first detectable ...

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13.9K Views.  Saint Louis University. Measuring left ventricular pressure (LV) in embryonic and neonatal mice is described. Pressure is measured by inserting a needle connected to a fluid-filled transducer into the LV under ultrasound guidance. Care must be taken to maintain normal cardiac function during the experimental protocol.

Video: Measuring Left Ventricular Pressure in Late Embryonic and Neonatal Mice

Patient-specific Modeling of the Heart: Estimation of Ventricular Fiber Orientations

Patient-specific simulations of heart (dys)function aimed at personalizing cardiac therapy are hampered by the absence of in vivo imaging technology for clinically acquiring myocardial fiber orientations. The objective of this project was to develop a methodology to estimate cardiac fiber orientations from in vivo images of patient heart geometries. An accurate representation of ventricular geometry and fiber orientations was reconstructed, respectively, from high-resolution ex vivo structural magnetic resonance (MR) and diffusion tensor (DT) MR images of a normal human heart, referred to as the atlas. Ventricular geometry of a patient heart was extracted, via semiautomatic segmentation, from an in vivo computed tomography (CT) image. Using image transformation algorithms, the atlas ventricular geometry was deformed to match that of the patient. Finally, the deformation field was applied to the atlas fiber orientations to obtain an estimate of patient fiber orientations. The accuracy of the fiber estimates was assessed using six normal and three failing canine hearts. The mean absolute difference between inclination angles of acquired and estimated fiber orientations was 15.4 &deg;. Computational simulations of ventricular activation maps and pseudo-ECGs in sinus rhythm and ventricular tachycardia indicated that there are no significant differences between estimated and acquired fiber orientations at a clinically observable level.The new insights obtained from the project will pave the way for the development of patient-specific models of the heart that can aid physicians in personalized diagnosis and decisions regarding electrophysiological interventions.

A methodology to estimate ventricular fiber orientations from in vivo images of patient heart geometries for personalized modeling is described. Validation of the methodology performed using normal and failing canine hearts demonstrate that that there are no significant differences between estimated and acquired fiber orientations at a clinically observable level.

Patient-specific  simulations of heart (dys)function aimed at personalizing cardiac therapy are hampered  by the absence of in vivo imaging technology for ...

Neonatal Cardiac Scaffolds: Novel Matrices for Regenerative Studies

The only definitive therapy for end stage heart failure is orthotopic heart transplantation. Each year, it is estimated that more than 100,000 donor hearts are needed for cardiac transplantation procedures in the United States1-2. Due to the limited numbers of donors, only approximately 2,400 transplants are performed each year in the U.S.2. Numerous approaches, from cell therapy studies to implantation of mechanical assist devices, have been undertaken, either alone or in combination, in an attempt to coax the heart to repair itself or to rest the failing heart3. In spite of these efforts, ventricular assist devices are still largely used for the purpose of bridging to transplantation and the utility of cell therapies, while they hold some curative promise, is still limited to clinical trials. Additionally, direct xenotransplantation has been attempted but success has been limited due to immune rejection. Clearly, another strategy is required to produce additional organs for transplantation and, ideally, these organs would be autologous so as to avoid the complications associated with rejection and lifetime immunosuppression. Decellularization is a process of removing resident cells from tissues to expose the native extracellular matrix (ECM) or scaffold. Perfusion decellularization offers complete preservation of the three dimensional structure of the tissue, while leaving the bulk of the mechanical properties of the tissue intact4. These scaffolds can be utilized for repopulation with healthy cells to generate research models and, possibly, much needed organs for transplantation. We have exposed the scaffolds from neonatal mice (P3), known to retain remarkable cardiac regenerative capabilities,5-8 to detergent mediated decellularization and we repopulated these scaffolds with murine cardiac cells. These studies support the feasibility of engineering a neonatal heart construct. They further allow for the investigation as to whether the ECM of early postnatal hearts may harbor cues that will result in improved recellularization strategies.

In these studies, we provide methodology for novel, neonatal, murine cardiac scaffolds for use in regenerative studies.

The only definitive therapy for end stage heart failure is orthotopic heart transplantation.  Each year, it is estimated that more than 100,000 donor ...

Measuring and Modeling Contractile Drying in Human Stratum Corneum

Stratum corneum (SC) is the most superficial skin layer. Its contact with the external environment means that this tissue layer is subjected to both cleansing agents and daily variations in ambient moisture; both of which can alter the water content of the tissue. Reductions in water content from severe barrier dysfunction or low humidity environments can alter SC stiffness and cause a build-up of drying stresses. In extreme conditions, these factors can cause mechanical rupture of the tissue. We have established a high throughput method of quantifying dynamic changes in the mechanical properties of SC upon drying. This technique can be employed to quantify changes in the drying behavior and mechanical properties of SC with cosmetic cleanser and moisturizer treatments. This is achieved by measuring dynamic variations in spatially resolved in-plane drying displacements of circular tissue samples adhered to an elastomer substrate. In-plane radial displacements acquired during drying are azimuthally averaged and fitted with a profile based on a linear elastic contractility model. Dynamic changes in drying stress and SC elastic modulus can then be extracted from the fitted model profiles.

This article describes a method of quantifying the dynamic drying behavior and mechanical properties of stratum corneum by measuring spatially resolved in-plane drying displacements of circular tissue samples adhered to an elastomer substrate. This technique can be used to measure how different chemical treatments alter drying and tissue mechanical properties.

Stratum corneum (SC) is the most superficial skin layer. Its contact with the external environment means that this tissue layer is subjected to both ...

Ammonia Synthesis at Low Pressure

Ammonia can be synthesized at low pressure by the use of an ammonia selective absorbent. The process can be driven with wind energy, available locally in areas requiring ammonia for synthetic fertilizer. Such wind energy is often called &#34;stranded,&#34; because it is only available far from population centers where it can be directly used.
In the proposed low pressure process, nitrogen is made from air using pressure swing absorption, and hydrogen is produced by electrolysis of water. While these gases can react at approximately 400 &#176;C in the presence of a promoted conventional catalyst, the conversion is often limited by the reverse reaction, which makes this reaction only feasible at high pressures. This limitation can be removed by absorption on an ammine-like calcium or magnesium chloride. Such alkaline metal halides can effectively remove ammonia, thus suppressing the equilibrium constraints of the reaction. In the proposed absorption-enhanced ammonia synthesis process, the rate of reaction may then be controlled not by the chemical kinetics nor the absorption rates, but by the rate of the recycle of unreacted gases. The results compare favorably with ammonia made from a conventional small scale Haber-Bosch process.

Ammonia can be synthesized at low pressure by using a conventional catalyst and an ammonia selective absorbent.

Ammonia can be synthesized at low pressure by the use of an ammonia selective absorbent. The process can be driven with wind energy, available locally in ...

Anatomically Realistic Neonatal Heart Model for Use in Neonatal Patient Simulators

Neonatal patient simulators (NPS) are artificial patient surrogates used in the context of medical simulation training. Neonatologists and nursing staff practice clinical interventions such as chest compressions to ensure patient survival in the case of bradycardia or cardiac arrest. The simulators used currently are of low physical fidelity and therefore cannot provide qualitative insight into the procedure of chest compressions. The embedding of an anatomically realistic heart model in future simulators enables the detection of cardiac output generated during chest compressions; this can provide clinicians with an output parameter, which can deepen the understanding of the effect of the compressions in relation to the amount of blood flow generated. Before this monitoring can be achieved, an anatomically realistic heart model must be created containing: two atria, two ventricles, four heart valves, pulmonary veins and arteries, and systemic veins and arteries. This protocol describes the procedure for creating such a functional artificial neonatal heart model by utilizing a combination of magnetic resonance imaging (MRI), 3D printing, and casting in the form of cold injection molding. Using this method with flexible 3D printed inner molds in the injection molding process, an anatomically realistic heart model can be obtained.

This protocol describes a procedure for creating functional artificial neonatal heart models by utilizing a combination of magnetic resonance imaging, 3D printing, and injection molding. The purpose of these models is for integration into the next generation of neonatal patient simulators and as a tool for physiological and anatomical studies.

Neonatal patient simulators (NPS) are artificial patient surrogates used in the context of medical simulation training. Neonatologists and nursing staff ...

Mouse Model of Pressure Ulcers After Spinal Cord Injury

Pressure ulcers (PUs) are common debilitating complications of traumatic spinal cord injury (SCI) and tend to occur in soft tissues around bony prominences. There is, however, little known about the impact of SCI on skin wound healing in the context of animal models in controlled experimental settings. In this study, a simple, non-invasive, reproducible and clinically relevant mouse model of PUs in the context of complete SCI is presented. Adult male mice (Balb/c, 10 weeks old) were shaved and depilated. Post-depilation (24 h), mice were subjected to laminectomy followed by complete spinal cord transection (T9-T10 vertebrae). Immediately after, a skin fold on the back of the mice was lifted and sandwiched between two magnetic discs held in place for next 12 h, thus creating an ischemic area that developed into a PU over the following days. The wounded areas demonstrated tissue edema and epidermal disappearance by day 3 post-magnet application. PUs spontaneously developed and healed. Healing was, however, slower in the SCI mice compared to control non-SCI mice when the wound was created below the level of SCI. Conversely, no difference in healing was seen between SCI and control non-SCI mice when the wound was created above the level of SCI. This model is a potentially useful tool to study the dynamics of skin PU development and healing after SCI, as well as to test therapeutic approaches that may help heal such wounds.

Here, we describe a simple method to induce clinically relevant skin pressure ulcers (PUs) in a mouse model of spinal cord injury (SCI). This model can be used in pre-clinical studies to screen for different therapeutics for healing PUs in SCI patients.

Pressure ulcers (PUs) are common debilitating complications of traumatic spinal cord injury (SCI) and tend to occur in soft tissues around bony ...

Measuring Global Cellular Matrix Metalloproteinase and Metabolic Activity in 3D Hydrogels

Three-dimensional (3D) cell culture systems often more closely recapitulate in vivo cellular responses and functions than traditional two-dimensional (2D) culture systems. However, measurement of cell function in 3D culture is often more challenging. Many biological assays require retrieval of cellular material which can be difficult in 3D cultures. One way to address this challenge is to develop new materials that enable measurement of cell function within the material. Here, a method is presented for measurement of cellular matrix metalloproteinase (MMP) activity in 3D hydrogels in a 96-well format. In this system, a poly(ethylene glycol) (PEG) hydrogel is functionalized with a fluorogenic MMP cleavable sensor. Cellular MMP activity is proportional to fluorescence intensity and can be measured with a standard microplate reader. Miniaturization of this assay to a 96-well format reduced the time required for experimental set up by 50% and reagent usage by 80% per condition as compared to the previous 24-well version of the assay. This assay is also compatible with other measurements of cellular function. For example, a metabolic activity assay is demonstrated here, which can be conducted simultaneously with MMP activity measurements within the same hydrogel. The assay is demonstrated with human melanoma cells encapsulated across a range of cell seeding densities to determine the appropriate encapsulation density for the working range of the assay. After 24 h of cell encapsulation, MMP and metabolic activity readouts were proportional to cell seeding density. While the assay is demonstrated here with one fluorogenic degradable substrate, the assay and methodology could be adapted for a wide variety of hydrogel systems and other fluorescent sensors. Such an assay provides a practical, efficient and easily accessible 3D culturing platform for a wide variety of applications.

Here, a protocol is presented for encapsulating and culturing cells in poly(ethylene glycol) (PEG) hydrogels functionalized with a fluorogenic matrix metalloproteinase (MMP)-degradable peptide. Cellular MMP and metabolic activity are measured directly from the hydrogel cultures using a standard microplate reader.

Three-dimensional (3D) cell culture systems often more closely recapitulate in vivo cellular responses and functions than traditional two-dimensional (2D) ...

Methods for In Vivo Biomechanical Testing on Brachial Plexus in Neonatal Piglets

Neonatal brachial plexus palsy (NBPP) is a stretch injury that occurs during the birthing process in nerve complexes located in the neck and shoulder regions, collectively referred to as the brachial plexus (BP). Despite recent advances in obstetrical care, the problem of NBPP continues to be a global health burden with an incidence of 1.5 cases per 1,000 live births. More severe types of this injury can cause permanent paralysis of the arm from the shoulder down. Prevention and treatment of NBPP warrants an understanding of the biomechanical and physiological responses of newborn BP nerves when subjected to stretch. Current knowledge of the newborn BP is extrapolated from adult animal or cadaveric BP tissue instead of in vivo neonatal BP tissue. This study describes an in vivo mechanical testing device and procedure to conduct in vivo biomechanical testing in neonatal piglets. The device consists of a clamp, actuator, load cell, and camera system that apply and monitor in vivo strains and loads until failure. The camera system also allows monitoring of the failure location during rupture. Overall, the presented method allows for a detailed biomechanical characterization of neonatal BP when subjected to stretch.

Methods for In Vivo Biomechanical Testing on Brachial Plexus in Neonatal Piglets

Presented here are methods to perform in vivo biomechanical testing on brachial plexus in a neonatal piglet model.

Neonatal brachial plexus palsy (NBPP) is a stretch injury that occurs during the birthing process in nerve complexes located in the neck and shoulder ...

Optical Imaging of Isolated Murine Ventricular Myocytes

The ability to isolate adult cardiac myocytes has permitted researchers to study a variety of cardiac pathologies at the single cell level. While advances in calcium sensitive dyes have permitted the robust optical recording of single cell calcium dynamics, recording of robust transmembrane optical voltage signals has remained difficult. Arguably, this is because of the low single to noise ratio, phototoxicity, and photobleaching of traditional potentiometric dyes. Therefore, single cell voltage measurements have long been confined to the patch clamp technique which while the gold standard, is technically demanding and low throughput. However, with the development of novel potentiometric dyes, large, fast optical responses to changes in voltage can be obtained with little to no phototoxicity and photobleaching. This protocol describes in detail how to isolate adult murine myocytes which can be used for cellular shortening, calcium, and optical voltage measurements. Specifically, the protocol describes how to use a ratiometric calcium dye, a single-excitation calcium dye, and a single excitation voltage dye. This approach can be used to assess the cardiotoxicity and arrhythmogenicity of various chemical agents. While phototoxicity is still an issue at the single cell level, methodology is discussed on how to reduce it.

We present the methodology for the isolation of murine myocytes and how to obtain voltage or calcium traces simultaneously with sarcomere shortening traces using fluorescence photometry with simultaneous digital cell geometry measurements.

The ability to isolate adult cardiac myocytes has permitted researchers to study a variety of cardiac pathologies at the single cell level. While advances ...

Patterning the Geometry of Human Embryonic Stem Cell Colonies on Compliant Substrates to Control Tissue-Level Mechanics

Human embryonic stem cells demonstrate a unique ability to respond to morphogens in vitro by self-organizing patterns of cell fate specification that correspond to primary germ layer formation during embryogenesis. Thus, these cells represent a powerful tool with which to examine the mechanisms that drive early human development. We have developed a method to culture human embryonic stem cells in confined colonies on compliant substrates that provides control over both the geometry of the colonies and their mechanical environment in order to recapitulate the physical parameters that underlie embryogenesis. The key feature of this method is the ability to generate polyacrylamide hydrogels with defined patterns of extracellular matrix ligand at the surface to promote cell attachment. This is achieved by fabricating stencils with the desired geometric patterns, using these stencils to create patterns of extracellular matrix ligand on glass coverslips, and transferring these patterns to polyacrylamide hydrogels during polymerization. This method is also compatible with traction force microscopy, allowing the user to measure and map the distribution of cell-generated forces within the confined colonies. In combination with standard biochemical assays, these measurements can be used to examine the role mechanical cues play in fate specification and morphogenesis during early human development.

Extracellular matrix ligands can be patterned onto polyacrylamide hydrogels to enable the culture of human embryonic stem cells in confined colonies on compliant substrates. This method can be combined with traction force microscopy and biochemical assays to examine the interplay between tissue geometry, cell-generated forces, and fate specification.

Human embryonic stem cells demonstrate a unique ability to respond to morphogens in vitro by self-organizing patterns of cell fate specification that ...

Measuring Left Ventricular Pressure in Late Embryonic and Neonatal Mice

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Anatomically Realistic Neonatal Heart Model for Use in Neonatal Patient Simulators

Mouse Model of Pressure Ulcers After Spinal Cord Injury

Measuring Global Cellular Matrix Metalloproteinase and Metabolic Activity in 3D Hydrogels

Methods for In Vivo Biomechanical Testing on Brachial Plexus in Neonatal Piglets

Optical Imaging of Isolated Murine Ventricular Myocytes

Patterning the Geometry of Human Embryonic Stem Cell Colonies on Compliant Substrates to Control Tissue-Level Mechanics