This research was performed within the Dutch framework of IMPULS perinatology. The authors would like to thank the Radboud UMCN Museum for Anatomy and Pathology and the M&#225;xima Medical Centre Veldhoven for providing the neonatal MRI scans used for this work. The authors further would like to thank Jasper Sterk, Sanne van der Linden, Frederique de Jongh, Pleun Alkemade, and the D.search lab at the faculty of Industrial Design for their significant contributions to the development of this research. Lastly the authors would like to thank Rohan Joshi for his proof reading of the manuscript.

All institutional approvals were obtained before patient imaging.
1. Image Acquisition and Segmentation
<ol>
	<li>Acquire a thoracic MRI scan of a neonate in Digital Imaging and Communications in Medicine (DICOM) format. Capture every slice of the scan in the ventricular diastolic stage of the heart cycle or obtain a thoracic MRI from an autopsy. 
	NOTE: A visibly clear definition of the heart muscle, as well as atria and ventricles, is essential.</li>
	<li>Using processing software (see Table of Materials) import the DICOM file of the thoracic MRI. Using the &#39;Editing Masks&#39; menu item, select the area of the heart muscle on each MRI slice where the heart is present. The atria and ventricles, in this case, can be covered as well.</li>
	<li>Create a new sketch layer and separately segment the two atria and two ventricles in the same manner as the selection for the heart muscle. Do not segment the valves present between the atria and ventricles, and between the ventricles and arteries.</li>
	<li>Render the muscle and chambers into separate 3D representations using the &#39;Calculate 3D&#39; menu item and export them as five STereoLithography (.stl) files using the optimal resolution settings by using the &#39;STL+&#39; menu item.</li>
	<li>Load the.stl files into the CAD software (see Table of Materials). Use the fix wizard menu item to repair the.stl files for overlapping triangles and bad edges. Save the.stl files again. 
	NOTE: If no heart MRI is available, consider using the heart model used in this protocol. This file also contains separate heart valve models. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/56710/Cardiac_model_parts.zip" target="_blank">Please click here to download the files</a>.</li>
</ol>
2. Processing and Mold Printing
<ol>
	<li>Load the set of atria and ventricles into a computer-aided design software (see Table of Materials).&#160;<a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/56710/Cardiac_model_parts.zip" target="_blank">Please click here to download the files</a>.
	<ol>
		<li>Determine the position of the aortic, pulmonary, mitral, and tricuspid valves using the original MRI (Figure 1).</li>
	</ol>
	</li>
	<li>Add the positive and negative mold halves of each valve to their respective position in the loaded set of atria and ventricles by dragging the valve file (obtained through the link above) into the current file activating the &#39;insert part&#39; function. Indicate the position of placement by clicking the location of the surface of the atria or ventricles.
	<ol>
		<li>Extrude the base of the positive and negative valve using &#39;features tab &#62; extrude boss/base&#39; to protrude into their respective chambers and merge the valve parts to their respective chamber. 
		NOTE: The mitral valve consists of two semilunar parts, while the tricuspid, aortic, and pulmonary valves consist of three.</li>
	</ol>
	</li>
	<li>Add the pulmonary and aortic valve file to their respective ventricle location using the procedure described in step 2.2. From the top of these valves, sketch two arching cylinders of 5 mm diameter by clicking the &#39;sketch tab &#62; circle&#39; following a sketched arched line using the &#39;features tab &#62; sweep boss/base&#39; until both circular cylinder surfaces reach the horizontal position. Merge the valve parts to their respective ventricles and arteries.</li>
	<li>From the base of each of the four chambers, as well as the two arched cylinders, draw vertical cylinders of 5 mm diameter by clicking the &#39;sketch tab &#62; circle&#39; item and extrude them to 40 mm in length by clicking the &#39;features tab &#62; extrude boss/base&#39; item. Let each cylinder protrude into their respective chamber.
	<ol>
		<li>To ensure the chambers positioning when assembling the six inner parts in the mold, add differential notches to the six cylinders (Figure 2) by sketching semicircles on top of the cylinders: click the &#39;sketch tab &#62; sketch circle&#39; menu item and use the &#39;feature tab &#62; cut/extrude&#39; menu item to create different depth indentations.
		<ol>
			<li>Subtract their shapes from the chambers and arteries by selecting the solid body of the chamber and artery, right clicking, and pressing the &#39;combine&#39; function after which the subtract setting can be selected. Do not merge these parts. Save all chambers and arteries separately.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Import the heart muscle model. Offset the six-cylinder base sketches by starting a new sketch and selecting all cylinder base sketches by holding down the &#39;shift&#39; key. Then, select the &#39;sketch tab &#62; convert entities&#39; menu item. Select the &#39;sketch tab &#62; offset entities&#39; menu item to offset the sketches by 2 mm.
	<ol>
		<li>Extrude and merge these sketches by clicking the &#39;features tab &#62; extrude boss/bass&#39; menu item with the heart muscle model; repeat for the arching cylinders. Merge these cylinders with the heart muscle model by clicking the &#39;features tab &#62; extrude boss/bass&#39; menu item. 
		NOTE: Ensure the heart muscle model opposite of the atria is over 2 mm in distance (Figure 1). Otherwise the wall will rupture when removing the inner molds.</li>
	</ol>
	</li>
	<li>Model a cube from the base of the six cylinders down by first placing a reference plane by clicking the &#39;features tab &#62; reference geometry &#62; plane&#39;. After this, click the &#39;sketch tab &#62; square&#39; menu item and sketch a square with a length and width that is 4 mm wider than the widest part of the heart muscle model.
	<ol>
		<li>Extrude this downwards with a thickness of 8 mm by clicking the &#39;features tab &#62; extrude boss/base&#39; menu item, and merge this to the base of the six cylinders marking the &#39;merge parts&#39; menu item. On the four corners of the base, add 4 mm cubes using the same method.</li>
	</ol>
	</li>
	<li>Using the square base as a sketch, extrude it to cover the entire heart model and subtract all other parts from this. Split the top part of the leftover rectangle at the widest part of the heart model. First place a reference plane at the desired height using the &#39;features tab &#62; reference geometry &#62; plane. After this, use the menu item &#39;insert &#62; molds &#62; split&#39; to select the surface over which the split has to take place and the object requiring splitting.
	<ol>
		<li>Split the leftover rectangle again at the most convenient mold release position using the same method described in step 2.7 yet in a vertical position. Sketch 4 mm cubic sockets in the corners of the longitudinal parts of the mold and add 4 mm cubes to the corners of the top cover using the &#39;sketch tab &#62; square&#39; and &#39;features tab &#62; extrude boss/base&#39; menu items.</li>
	</ol>
	</li>
	<li>Sketch 50 circles of 1 mm in diameter covering the top of the entire outer mold model and cut-extrude these through all outer molds. Also, extrude several 1 mm cylinders on the side of the top cover at the widest locations of the heart muscle model. Cut-extrude a single 8 mm injection hole from the top cover.
	<ol>
		<li>Save all four outer mold parts separately. 
		CAUTION: In total, there should be ten mold components: the base of the mold, two outer mold side panels, one outer mold top cover, two inner mold atria with valve attachments, two inner mold ventricles with valve attachments, and one of each aortic and pulmonary inner mold artery with valve attachments.</li>
	</ol>
	</li>
	<li>Use a jetting printer for printing with rigid and rubber-like photopolymer materials installed 
	(see Table of Materials). When placing the parts for printing on the print bed, ensure the valve negatives are all printed facing upwards (vertically) (Figure 3).
	<ol>
		<li>Select the print settings to glossy. For the four chambers as well as the pulmonary and aortic mold attachments, select the flexible S95 material; for the other four mold parts, select the rigid print material.</li>
	</ol>
	</li>
	<li>After printing the mold parts, remove the support material built up during printing by waterjet (see Table of Materials). After cleaning the mold parts, place the parts in a 5% sodium hydroxide solution for 24 h. After removing the parts from the solution, rinse them using cold water and leave to dry for 48 h before casting.</li>
</ol>
3. Cold Injection Molding and Finishing
<ol>
	<li>Spray all surfaces of all mold parts with a release agent (see Table of Materials), except the valves, and wipe clean with tissue paper. Leave to dry for 15 min.
	<ol>
		<li>Close the base of the mold and two side panels, and place on top of two spacers, so the base of the mold is not in direct contact with the table surface. Prepare the silicone by inserting a silicone cartridge into the manual dispensing gun (see Table of Materials).</li>
	</ol>
	</li>
	<li>Add 5 mL of silicone squeezed from the dispensing gun into a measuring cup and mix using a toothpick. Using a toothpick, apply a generous amount of melted silicone to the negative and positive side of the right atria and ventricle valves. Make sure there are no air bubbles entrapped in the silicone (Figure 4).
	<ol>
		<li>Connect the two chambers at the right valve angle, and push them onto their respective cylinders of the base mold. Repeat this for the left side. Finally, attach the pulmonary and aortic arched cylinders similarly. Leave these valves to solidify for 2 min, then attach the top part of the mold.</li>
	</ol>
	</li>
	<li>Attach a static mixer to the cartridge, squeeze until the silicone is exiting the nozzle, then release the pressure. Position the entire mold on two spacers (Figure 5), insert the gun into the 8-mm injection molding socket, and squeeze with low pressure over the course of 3 min until all the air vents show signs of silicone overflow.
	<ol>
		<li>Stop injecting the silicone at this point, remove the mixer, and place the mold onto the table surface so that all bottom air vents are sealed, and no more silicone can flow from the bottom of the mold. Leave the silicone to solidify for 30 min.</li>
	</ol>
	</li>
	<li>Open the top part of the mold by prying and lifting a metal spacer into the crack between the top and lower part of the mold. Remove the side parts of the mold using the same method, removing one side at a time. 
	NOTE: Make sure not to puncture the heart wall when inserting the spacer.
	<ol>
		<li>Detect any air bubbles at the heart exterior after releasing the three outer mold components (Figure 6). Use a scalpel to pierce the bubble and fill it with a small amount silicone using a toothpick, then leave to cure for another 30 min.</li>
	</ol>
	</li>
	<li>Use compressed air (see Table of Materials) to blow the heart model off the base of the mold leaving the six inner molds in the heart model. Make sure to firmly enclose the heart model with one hand to prevent air rupturing the heart wall.
	<ol>
		<li>Use a syringe with water to fill and pressurize the left and right ventricles to release the inner molds. After this use a Magill forceps (see Table of Materials) to grab and pull out these inner two mold parts. Repeat this process for the pulmonary and aortic arteries and lastly for removing the left and right atria inner molds. 
		NOTE: Make sure the positioning of the forceps does not compress the valve segment when clamp pressure is applied; it will destroy the printed valve.</li>
	</ol>
	</li>
	<li>Bind the two tubes leading directly downwards from the ventricles at the base of the heart model using tie wraps and remove access air vent strings by plucking them at the heart wall surface.</li>
</ol>

<ol>
	<li>Wyllie, J., Bruinenberg, J., Roehr, C. C., R&#252;diger, M., Trevisanuto, D., Urlesberger, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=European+resuscitation+council+guidelines+for+resuscitation+2015.">European resuscitation council guidelines for resuscitation 2015.</a> Resuscitation. 95, 249-263 (2015).</li><li>Sawyer, T., Strandjord, T. P., Johnson, K., Low, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Neonatal+airway+simulators,+how+good+are+they?+A+comparative+study+of+physical+and+functional+fidelity.">Neonatal airway simulators, how good are they? A comparative study of physical and functional fidelity.</a> J. Perinatol. 36 (2), 151-156 (2015).</li><li>Yao, R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Three-dimensional+printing:+review+of+application+in+medicine+and+hepatic+surgery.">Three-dimensional printing: review of application in medicine and hepatic surgery.</a> Cancer Biol. Med. 13 (4), 443-451 (2016).</li><li>Chua, C. K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rapid+prototyping+assisted+surgery+planning.">Rapid prototyping assisted surgery planning.</a> Int. J. Adv. Manuf. Tech. 14 (9), 624-630 (1998).</li><li>Gibson, I., et al. <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+of+rapid+prototyping+to+assist+medical+applications.">The use of rapid prototyping to assist medical applications.</a> Rapid Prototyping J. 12 (1), 53-58 (2006).</li><li>Cai, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Application+of+3D+printing+in+orthopedics:+status+quo+and+opportunities+in+China.">Application of 3D printing in orthopedics: status quo and opportunities in China.</a> Ann. Transl. Med. 3 (Suppl 1), S12 (2015).</li><li>Thielen, M. W. H., Delbressine, F. L. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rib+cage+recreation:+towards+realistic+neonatal+manikin+construction+using+MRI+scanning+and+3D+printing.">Rib cage recreation: towards realistic neonatal manikin construction using MRI scanning and 3D printing.</a> FASE. , 41-44 (2016).</li><li>Thielen, M., Joshi, R., Delbressine, F., Bambang Oetomo, S., Feijs, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+innovative+design+for+cardiopulmonary+resuscitation+manikins+based+on+a+human-like+thorax+and+embedded+flow+sensors.">An innovative design for cardiopulmonary resuscitation manikins based on a human-like thorax and embedded flow sensors.</a> JOEIM. 231 (3), 243-249 (2017).</li><li>Cohrs, N. C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+soft+Total+Artificial+Heart+-+First+Concept+Evaluation+on+a+Hybrid+Mock+Circulation.">A soft Total Artificial Heart - First Concept Evaluation on a Hybrid Mock Circulation.</a> Artif. Organs. , (2017).</li><li>Sparks, J. L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Use+of+silicone+materials+to+simulate+tissue+biomechanics+as+related+to+deep+tissue+injury.">Use of silicone materials to simulate tissue biomechanics as related to deep tissue injury.</a> Adv. Skin Wound Care. 28 (2), 59-68 (2015).</li><li>Van der Horst, A., Geven, M. C., Rutten, M. C., Pijls, N. H., Nvan de Vosse, F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Thermal+anemometric+assessment+of+coronary+flow+reserve+with+a+pressure-sensing+guide+wire:+An+in+vitro+evaluation.">Thermal anemometric assessment of coronary flow reserve with a pressure-sensing guide wire: An in vitro evaluation.</a> Med. Eng. Phys. 33 (6), 684-691 (2011).</li><li>Miriyev, A., Stack, K., Lipson, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Soft+material+for+soft+actuators.">Soft material for soft actuators.</a> Nature comm. 8 (596), (2017).</li></ol>

The authors declare no potential conflicts of interest concerning research, authorship, and publication of this article. This research received no specific grant from any funding agency in public, commercial, or not-for-profit sectors.

For the model developed in this study, we identified that injection molding over a 3-min period is required to prevent air entering the cast (Figure 5, Figure 6). To ensure that silicone reaches the narrow spaces of the valves, &#34;pre-casting&#34; or &#34;coating&#34; of the valve areas in the mold is essential. Since the inner molds shaping the heart chambers have to exit the final silicone cast through 5 mm openings, multi-material 3D printing for molds is n...

Every year millions of neonates are admitted to neonatal intensive care units (NICU). In NICUs, most emergencies relate to problems in the airway, breathing, and circulation (ABC) and require interventions such as chest compressions. NPS offer a valuable teaching and training tool to practice such interventions. For some NPS, embedded sensors can detect whether performance meets the recommended clinical guidelines1 for depth and speed of chest compressions. The adherence to guidelines can be used to calculate and quantify performance, and in this regard, such state of the art NPS can be viewed as a tangible and white box metric for evaluating performance.
Adherence to the recommended guidelines aims at improving patient physiology. For example, chest compressions are delivered with the aim of generating adequate blood flow in the circulatory system. Current high fidelity NPS (e.g., PremieAnne (Laerdal, Stavanger, Norway) and Paul (SIMCharacters, Vienna, Austria)), do not contain any sensors to measure physiological parameters such as blood flow during training as they lack an integrated heart to generate this physiological parameter. Efficacy of chest compressions in current NPS can therefore not be assessed at a physiological level. For NPS to enable physiological assessment of chest compressions, an anatomically realistic artificial heart has to be integrated into the NPS. Furthermore, research2 shows that an increase in physical anatomical fidelity may lead to an increase in functional fidelity of NPS. Integrating a physically high-fidelity organ system would benefit both the functional fidelity of training and enable physiological performance assessment.
A substantial increase in the fidelity of NPS can be achieved through 3D printing. In medicine, 3D imaging and printing are mostly used for surgical preparation and creation of implants3,4,5. For example, in the field of surgical simulation, organs are produced to train surgeons on performing surgical procedures6. The possibilities of 3D printing have not yet been extensively applied in NPS. The combination of 3D imaging and 3D printing opens the possibility for NPS to reach a higher level of physical fidelity. The replication of sophisticated, flexible, neonatal organs such as the heart becomes possible due to the ever-broadening range of techniques and materials used for 3D printing7.
In this paper, we detail a protocol for creating a functional, artificial neonatal heart using a combination of MRI, 3D printing, and cold injection molding. The heart model in this paper includes two atria, two ventricles, four functional valves, and pulmonary and systemic arteries and veins all produced from a single silicone cast. The heart model can be filled with a liquid, equipped with sensors, and used as output parameter generator (i.e., blood pressure or cardiac output during chest compressions, and valve functionality).

<table>
	<tbody>
		<tr>
			<td>Ecoflex 5</td>
			<td>Smooth-on</td>
			<td></td>
			<td>Silicon casting material</td>
		</tr>
		<tr>
			<td>400ml Static mixers</td>
			<td>Smooth-on</td>
			<td></td>
			<td>Mixing tubes</td>
		</tr>
		<tr>
			<td>Manual dispensing gun</td>
			<td>Smooth-on</td>
			<td></td>
			<td>Used for injection molding</td>
		</tr>
		<tr>
			<td>5-56 PTFE spray</td>
			<td>CRC</td>
			<td></td>
			<td>Release agent for the molds</td>
		</tr>
		<tr>
			<td>Sodium-hydroxide</td>
			<td>N/A</td>
			<td></td>
			<td>This was purchased as caustic soda at the hardware store, in dry, 99% pure form. As it is widely available, there is no company specified</td>
		</tr>
		<tr>
			<td>VeroWhite</td>
			<td>Stratasys</td>
			<td></td>
			<td>The hard material used in the print</td>
		</tr>
		<tr>
			<td>TangoBlackPlus</td>
			<td>Stratasys</td>
			<td></td>
			<td>The rubber material used in the print</td>
		</tr>
		<tr>
			<td>Support Material</td>
			<td>Stratasys</td>
			<td></td>
			<td>The standard support material used by stratasys&#160;</td>
		</tr>
		<tr>
			<td>Magill Forceps</td>
			<td>GIMA</td>
			<td></td>
			<td>Infant size. This is for removing the inner molds</td>
		</tr>
		<tr>
			<td>Stratasys Connex 350</td>
			<td>Stratasys</td>
			<td></td>
			<td>&#160;If this machine is not owned, another option is to have the parts printed through a third party printing firm such as 3D-hubs to get the parts printed and shipped.</td>
		</tr>
		<tr>
			<td>Balco Powerblast (Water Jet)</td>
			<td>Stratasys</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Euro 8-24 Set P (Air Compressor)</td>
			<td>iSC</td>
			<td>4007292</td>
			<td></td>
		</tr>
		<tr>
			<td>Syringe with blunt needle</td>
			<td>N/A</td>
			<td></td>
			<td>A 20ml syringe with a 0.5mm diameter blunt needle.</td>
		</tr>
		<tr>
			<td>Mimics 17.0 software</td>
			<td>Materialise&#160;</td>
			<td></td>
			<td>This software was used to segment the heart model from the MRI. There are sevaral free MRI imaging software tools available such as InVesalius, or Osirix, although they may prove to provide less functionality.</td>
		</tr>
		<tr>
			<td>Magics 9.0 software</td>
			<td>Materialise&#160;</td>
			<td></td>
			<td>This was used to repair and smooth the .stl files generated by mimics. This smoothing can also&#160; be done in most other 3D modeling freeware.</td>
		</tr>
		<tr>
			<td>Solidworks</td>
			<td></td>
			<td></td>
			<td>Software used for editting the heart model. Most other freeware CAD software can be used to perform this stage of processing.</td>
		</tr>
	</tbody>
</table>

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 study details a method to create an anatomically realistic neonatal heart model combining MRI imaging, 3D printing, and cold injection molding. The ductus arteriosus as well as foramen ovale were not included in the heart model presented in this paper. The method described in this paper can also be applied to other internal organs, such as lungs, and rib cage structures. Rib cage structures require no molds and can be printed directly using flexible materials. In (<strong class="xfig...

Watch this Scientific Journal Video about Anatomically Realistic Neonatal Heart Model for Use in Neonatal Patient Simulators at JoVE.com

Anatomically Realistic Neonatal Heart Model for Use in Neonatal Patient Simulators

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 ...

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Universidad de Cartagena UDEC

Research

JoVE Journal

Bioengineering

5.9K Views.  Eindhoven University of Technology. 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.

Video: Anatomically Realistic Neonatal Heart Model for Use in Neonatal Patient Simulators

Multiparametric Optical Mapping of the Langendorff-perfused Rabbit Heart

Optical imaging and fluorescent probes have significantly advanced research methodology in the field of cardiac electrophysiology in ways that could not have been accomplished by other approaches1. With the use of the calcium- and voltage-sensitive dyes, optical mapping allows measurement of transmembrane action potentials and calcium transients with high spatial resolution without the physical contact with the tissue. This makes measurements of the cardiac electrical activity possible under many conditions where the use of electrodes is inconvenient or impossible1. For example, optical recordings provide accurate morphological changes of membrane potential during and immediately after stimulation and defibrillation, while conventional electrode techniques suffer from stimulus-induced artifacts during and after stimuli due to electrode polarization1. 
The Langendorff-perfused rabbit heart is one of the most studied models of human heart physiology and pathophysiology. Many types of arrhythmias observed clinically could be recapitulated in the rabbit heart model. It was shown that wave patterns in the rabbit heart during ventricular arrhythmias, determined by effective size of the heart and the wavelength of reentry, are very similar to that in the human heart2. It was also shown that critical aspects of excitation-contraction (EC) coupling in rabbit myocardium, such as the relative contribution of sarcoplasmic reticulum (SR), is very similar to human EC coupling3. Here we present the basic procedures of optical mapping experiments in Langendorff-perfused rabbit hearts, including the Langendorff perfusion system setup, the optical mapping systems setup, the isolation and cannulation of the heart, perfusion and dye-staining of the heart, excitation-contraction uncoupling, and collection of optical signals. These methods could be also applied to the heart from species other than rabbit with adjustments to flow rates, optics, solutions, etc.
Two optical mapping systems are described. The panoramic mapping system is used to map the entire epicardium of the rabbit heart4-7. This system provides a global view of the evolution of reentrant circuits during arrhythmogenesis and defibrillation, and has been used to study the mechanisms of arrhythmias and antiarrhythmia therapy8,9. The dual mapping system is used to map the action potential (AP) and calcium transient (CaT) simultaneously from the same field of view10-13. This approach has enhanced our understanding of the important role of calcium in the electrical alternans and the induction of arrhythmia14-16.

This article describes the basic procedures for conducting optical mapping experiments in the Langendorff-perfused rabbit heart using the panoramic imaging system, and the dual (voltage and calcium) imaging modality.

Optical imaging and fluorescent probes have significantly advanced research methodology in the field of cardiac electrophysiology in ways that could not ...

Optical Mapping of Action Potentials and Calcium Transients in the Mouse Heart

The mouse heart is a popular model for cardiovascular studies due to the existence of low cost technology for genetic engineering in this species. Cardiovascular physiological phenotyping of the mouse heart can be easily done using fluorescence imaging employing various probes for transmembrane potential (Vm), calcium transients (CaT), and other parameters. Excitation-contraction coupling is characterized by action potential and intracellular calcium dynamics; therefore, it is critically important to map both Vm and CaT simultaneously from the same location on the heart1-4. Simultaneous optical mapping from Langendorff perfused mouse hearts has the potential to elucidate mechanisms underlying heart failure, arrhythmias, metabolic disease, and other heart diseases. Visualization of activation, conduction velocity, action potential duration, and other parameters at a myriad of sites cannot be achieved from cellular level investigation but is well solved by optical mapping1,5,6. In this paper we present the instrumentation setup and experimental conditions for simultaneous optical mapping of Vm and CaT in mouse hearts with high spatio-temporal resolution using state-of-the-art CMOS imaging technology. Consistent optical recordings obtained with this method illustrate that simultaneous optical mapping of Langendorff perfused mouse hearts is both feasible and reliable.

This paper details the dissection procedure, instrumental setup, and experimental conditions during optical mapping of transmembrane potential (Vm) and intracellular calcium transient (CaT) in intact isolated Langendorff perfused mouse hearts.

The mouse heart is a popular model for cardiovascular studies due to the existence of low cost technology for genetic engineering in this species. ...

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.

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 ...

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 ...

An In Vitro Organ Culture Model of the Murine Intervertebral Disc

Intervertebral disc (IVD) degeneration is a significant contributor to low back pain. The IVD is a fibrocartilaginous joint that serves to transmit and dampen loads in the spine. The IVD consists of a proteoglycan-rich nucleus pulposus (NP) and a collagen-rich annulus fibrosis (AF) sandwiched by cartilaginous end-plates. Together with the adjacent vertebrae, the vertebrae-IVD structure forms a functional spine unit (FSU). These microstructures contain unique cell types as well as unique extracellular matrices. Whole organ culture of the FSU preserves the native extracellular matrix, cell differentiation phenotypes, and cellular-matrix interactions. Thus, organ culture techniques are particularly useful for investigating the complex biological mechanisms of the IVD. Here, we describe a high-throughput approach for culturing whole lumbar mouse FSUs that provides an ideal platform for studying disease mechanisms and therapies for the IVD. Furthermore, we describe several applications that utilize this organ culture method to conduct further studies including contrast-enhanced microCT imaging and three-dimensional high-resolution finite element modeling of the IVD.

An In Vitro Organ Culture Model of the Murine Intervertebral Disc

Whole organ culture of the intervertebral disc (IVD) preserves the native extracellular matrix, cell phenotypes, and cellular-matrix interactions. Here we describe an IVD culture system using mouse lumbar and caudal IVDs in their functional spinal units and several applications utilizing this system.

Intervertebral disc (IVD) degeneration is a significant contributor to low back pain. The IVD is a fibrocartilaginous joint that serves to transmit and ...

Creating a Structurally Realistic Finite Element Geometric Model of a Cardiomyocyte to Study the Role of Cellular Architecture in Cardiomyocyte Systems Biology

With the advent of three-dimensional (3D) imaging technologies such as electron tomography, serial-block-face scanning electron microscopy and confocal microscopy, the scientific community has unprecedented access to large datasets at sub-micrometer resolution that characterize the architectural remodeling that accompanies changes in cardiomyocyte function in health and disease. However, these datasets have been under-utilized for&#160;investigating the role of cellular architecture remodeling in cardiomyocyte function. The purpose of this protocol is to outline how to create an accurate finite element model of a cardiomyocyte using high resolution electron microscopy and confocal microscopy images. A detailed and accurate model of cellular architecture has significant potential to provide new insights into cardiomyocyte biology, more than experiments alone can garner. The power of this method lies in its ability to computationally fuse information from two disparate imaging modalities of cardiomyocyte ultrastructure to develop one unified and detailed model of the cardiomyocyte. This protocol outlines steps to integrate electron tomography and confocal microscopy images of adult male Wistar (name for a specific breed of albino rat) rat cardiomyocytes to develop a half-sarcomere finite element model of the cardiomyocyte. The procedure generates a 3D finite element model that contains an accurate, high-resolution depiction (on the order of ~35 nm) of the distribution of mitochondria, myofibrils and ryanodine receptor clusters that release the necessary calcium for cardiomyocyte contraction from the sarcoplasmic reticular network (SR) into the myofibril and cytosolic compartment. The model generated here as an illustration does not incorporate details of the transverse-tubule architecture or the sarcoplasmic reticular network and is therefore a minimal model of the cardiomyocyte. Nevertheless, the model can already be applied in simulation-based investigations into the role of cell structure in calcium signaling and mitochondrial bioenergetics, which is illustrated and discussed using two case studies that are presented following the detailed protocol.

This protocol outlines a novel method to create a spatially detailed finite element model of the intracellular architecture of cardiomyocytes from electron microscopy and confocal microscopy images. The power of this spatially detailed model is demonstrated using case studies in calcium signaling and bioenergetics.

With the advent of three-dimensional (3D) imaging technologies such as electron tomography, serial-block-face scanning electron microscopy and confocal ...

Recombinant Collagen I Peptide Microcarriers for Cell Expansion and Their Potential Use As Cell Delivery System in a Bioreactor Model

Tissue engineering is a promising field, focused on developing solutions for the increasing demand on tissues and organs regarding transplantation purposes. The process to generate such tissues is complex, and includes an appropriate combination of specific cell types, scaffolds, and physical or biochemical stimuli to guide cell growth and differentiation. Microcarriers represent an appealing tool to expand cells in a three-dimensional (3D) microenvironment, since they provide higher surface-to volume ratios and mimic more closely the in vivo situation compared to traditional two-dimensional methods. The vascular system, supplying oxygen and nutrients to the cells and ensuring waste removal, constitutes an important building block when generating engineered tissues. In fact, most constructs fail after being implanted due to lacking vascular support. In this study, we present a protocol for endothelial cell expansion on recombinant collagen-based microcarriers under dynamic conditions in spinner flask and bioreactors, and we explain how to determine in this setting cell viability and functionality. In addition, we propose a method for cell delivery for vascularization purposes without additional detachment steps necessary. Furthermore, we provide a strategy to evaluate the cell vascularization potential in a perfusion bioreactor on a decellularized&#160;biological matrix. We believe that the use of the presented methods could lead to the development of new cell-based therapies for a large range of tissue engineering applications in the clinical practice.

We propose a cell expansion protocol on macroporous microcarriers and their use as delivery system in a perfusion bioreactor to seed a decellularized tissue matrix. We also include different techniques to determine cell proliferation and viability of cells cultured on microcarriers. Furthermore, we demonstrate functionality of cells after bioreactor cultures.

Tissue engineering is a promising field, focused on developing solutions for the increasing demand on tissues and organs regarding transplantation ...

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 ...

Use of High-Throughput Automated Microbioreactor System for Production of Model IgG1 in CHO Cells

Automated microscale bioreactors (15 mL) can be a useful tool for cell culture engineers. They facilitate the simultaneous execution of a wide variety of experimental conditions while minimizing potential process variability. Applications of this approach include: clone screening, temperature and pH shifts,&#160;media and supplement optimization. Furthermore, the small reactor volumes are conducive to large Design of Experiments that investigate a wide range of conditions. This allows upstream processes to be significantly optimized before scale-up where experimentation is more limited in scope due to time and economic constraints. Automated microscale bioreactor systems offer various advantages over traditional small scale cell culture units, such as shake flasks or spinner flasks. However, during pilot scale process development&#160;significant care must be taken to ensure that these advantages are realized. When run with care, the system can enable high level automation, can be programmed to run DOE&#39;s with a higher number of variables&#160;and can reduce sampling time when integrated with a nutrient analyzer or cell counter. Integration of the expert-derived heuristics presented here, with current automated microscale bioreactor experiments can minimize common pitfalls that hinder meaningful results. In the extreme, failure to adhere to the principles laid out here can lead to equipment damage that requires expensive repairs. Furthermore, the microbioreactor systems have small culture volumes&#160;making characterization of cell culture conditions difficult. The number and amount of samples taken in-process in batch mode culture is limited as operating volumes cannot fall below 10 mL. This method will discuss the benefits and drawbacks of microscale bioreactor systems.

A detailed protocol for the concurrent operation of 48 parallel cell cultures under varied conditions in a microbioreactor system is presented. Cell culture process, harvest and subsequent antibody titer analysis are described.

Automated microscale bioreactors (15 mL) can be a useful tool for cell culture engineers. They facilitate the simultaneous execution of a wide variety of ...

Anatomically Realistic Neonatal Heart Model for Use in Neonatal Patient Simulators

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Chapters in this video

Multiparametric Optical Mapping of the Langendorff-perfused Rabbit Heart

Optical Mapping of Action Potentials and Calcium Transients in the Mouse Heart

Measuring Left Ventricular Pressure in Late Embryonic and Neonatal Mice

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

Neonatal Cardiac Scaffolds: Novel Matrices for Regenerative Studies

An In Vitro Organ Culture Model of the Murine Intervertebral Disc

Creating a Structurally Realistic Finite Element Geometric Model of a Cardiomyocyte to Study the Role of Cellular Architecture in Cardiomyocyte Systems Biology

Recombinant Collagen I Peptide Microcarriers for Cell Expansion and Their Potential Use As Cell Delivery System in a Bioreactor Model

Mouse Model of Pressure Ulcers After Spinal Cord Injury

Use of High-Throughput Automated Microbioreactor System for Production of Model IgG1 in CHO Cells