A seed grant from the University of Florida - College of Medicine is gratefully acknowledged. Graduate studies (Manuel Salinas) were supported through a minority opportunities in biomedical research programs - research initiative for scientific enhancement (MBRS-RISE) fellowship: NIH/NIGMS R25 GM061347. Financial support from the Wallace H. Coulter Foundation through Florida International University's, Biomedical Engineering Department is also gratefully acknowledged. Finally, the authors thank the following students for their assistance during various stages of the experimental process: Kamau Pier, Malachi Suttle, Kendall Armstrong and Abraham Alfonso.

1. Preparation
<ol>
	<li>Design and fabricate an assembly to accommodate a tri-leaflet valve geometry. This will at minimum include a valve holder to suture-in the valve leaflets&#160;and a tube to house the valve holder and surrounding accessories to secure the assembly onto the pulse duplicator system. In our case, we utilized a commercially available pulse duplicator system available from ViVitro Labs Inc. (Victoria, BC). Valve holder design as well as pre and post assembly configurations are depicted in Figure 1.</li>
	<li>The entire loop will need to be primed prior to usage. This involves two steps: i) cleansing of the entire loop system using soap solution and water, including replacement of any degraded tubes prior to use and ii) calibration of instruments connected to the loop, namely the pump being used, the flow probe, and the pressure transducers (generally measured at atrial, aortic and ventricular locations). Calibration can initially be performed using 1% saline solution and should be repeated prior to using blood-analog glycerin solution.</li>
</ol>
2. Native Aortic Valve Dissection
<ol>
	<li>Obtain 4 fresh pig hearts with the aorta intact from a USDA approved slaughter house (Institutional Animal Care and Use Committee (IACUC) approval may be required). In our case, our dissection protocol was approved by the IACUC at Florida International University (Protocol Approval Number: 11-020). Rinse the heart with deionized water and place it in a receptacle filled with the 1% antimycotic/antibiotic and sterile phosphate buffered saline (PBS) solution and transport on ice to the hydrodynamic testing laboratory.</li>
	<li>Place hearts in a dissecting pan and carefully remove the pericardium. Position the heart such that ventral side is facing you. Visually inspect and identify the four chambers of the heart and locate the aortic arch on the intact aorta.</li>
	<li>Separate the heart into two halves by cutting across horizontally at approximately 0.75 in&#160;below the annulus, i.e.&#160;the junction between the aorta and the left ventricle. Carefully isolate the intact aorta still attached to the left ventricular tissue segment.</li>
	<li>Examine the aortic valve located in the aortic root, the region between the ascending aorta and the lower annulus, ensuring that there is no damage or any signs of calcification.</li>
	<li>Split the aorta at ~1 in&#160;above the annulus and separate the left ventricular tissue segment below the annulus to isolate the aortic valve (Figure 2a).</li>
</ol>
3. Polymer and Native Valve Suturing Process
<ol>
	<li>Place the heart valve inside the valve holder such that the base of each valve aligns with the base of the post holder. Secure the valve in place at each post temporarily with a paper clip, but be careful not to damage the commissures or the cusps.</li>
	<li>Insert the suture in the needle. Begin suturing at the bottom of the valve holder by passing the needle through the first hole, from the outside to the inside such that the needle may be easily pulled from the bottom. In a looping fashion, start suturing the valve vertically up the posts of the valve holder.</li>
	<li>Progress with suturing (Figure 2b) along the circumference of the holder and secure with additional suture around the tips of the holder posts. Paper clips (Figure 2c) can be removed when the valve is completely secured using sutures to the 3 posts and at the circumference of the valve holder (Figures 2d and 2e).</li>
</ol>
4. Hydrodynamic Evaluation&#160;
Note: Actual protocol will vary depending on specific pulse duplicator system being used. All information caontained herein used the ViVitro Pulse Duplicator Sysytem (ViVitro Labs, Inc., Vancouver, BC).
<ol>
	<li>Bi-leaflet valve
	<ol>
		<li>Set heart rate of pulse duplicator system to 70 beats/min.</li>
		<li>Select a flow waveform to drive the pump (in the case of the ViVitro system the S35 waveform was chosen for all hydrodynamic tests). The specific waveform utilized in our experiments is illustrated by Lim et al.&#160;(2001)17.</li>
		<li>Turn on amplifier and piston pump. Warm up for 15 min.</li>
		<li>Place bi-leaflet valve (Figure 2f) in the aortic position.</li>
		<li>Smear vacuum grease on all junctions of the device where leaks could occur.</li>
		<li>Pour glycerin/saline liquid in the atrial compartment. Note that the pulsatile duplicator system runs on 2 L of liquid with: 35%/0.7 L glycerin and 65%/1.3 L of saline solution. The saline solution is prepared using common salt well-dissolved in deionized water at a concentration of 9 mg/ml (weight/volume).</li>
		<li>Turn on the flow transducer that has been placed in the aortic position.</li>
		<li>Calibrate the pump.</li>
		<li>Proceed with the flow transducer calibration followed by the pressure transducers. Similarly to the pump, simply follow the instructions given by the ViVitest software (ViVitro Labs Inc.) for each flow and pressure under the calibrate tab.</li>
		<li>Once calibration is complete, start the pump at a low rpm until the fluid fills the aortic compartment. Check for leaks. Use additional vacuum grease if necessary.</li>
		<li>Turn the two stop-cocks (aortic and ventricular transducers) to open position.</li>
		<li>Increase the rpm of the pump until the stroke volume reaches 80 ml/beat.</li>
		<li>Permit the system to run for 10 min until flow has stabilized. Flow stabilization can be verified by observing the flow and pressure waveforms displayed in the screen. Low to none variation between cycles is a good indicator of system stabilization.</li>
		<li>In the ViVitest software select acquire mode.</li>
		<li>Click on collect 10 cycles.</li>
		<li>From the analyze mode, click on table and save the file. Also save an image of the waveforms using the photo-snap option in ViVitest.</li>
	</ol>
	</li>
	<li>Native and Polymer valves
	<ol>
		<li>For polymer and animal valves, follow the same steps 3.1.1 -&#160;3.1.3 from the bi-leaflet valve instructions.</li>
		<li>Place the valve holder with the sutured valve inside the glass tube from the custom made assembly. Sandwich the tube with the top and bottom pieces and secure in-place with lateral screws and nuts.</li>
		<li>Place assembly between the aortic chamber and the original aortic valve holder.</li>
		<li>Continue with steps 3.1.5 -&#160;3.1.16 from the bi-leaflet valve instructions.</li>
	</ol>
	</li>
</ol>
5. Post Processing
<ol>
	<li>Flow and Pressure Waveforms
	<ol>
		<li>Average the data collected for each of the waveforms collected, i.e.&#160;aortic pressure (AP), ventricular pressure (VP), and flow rate (Q).</li>
		<li>For each group of valve (polymer, porcine native aortic valve and bi-leaflet), plot the corresponding AP, VP and Q versus time relationships on the same plot.</li>
		<li>For the AP, superimpose normal, native aortic valve18, and bi-leaflet prosthetic valve19 plots from the literature for validation purposes.</li>
	</ol>
	</li>
	<li>Hydrodynamic metrics
	<ol>
		<li>For each valve tested, the following hydrodynamic metrics should be computed: a) Forward flow pressure drop and maximum transvalvular pressure (TVP), b) the aortic root mean square (RMS) forward flow rate, c) aortic forward flow, closing, leakage and total regurgitant volume, d) valve end orifice area (EOA), e) transaortic forward flow, closing, leakage and total energy losses.
		<ol>
			<li>Forward flow pressure drop is computed from TVP readings and can be categorized into 3 time intervals, P: interval that starts and ends with 0 TVP, F: interval with forward flow and H: interval starting with 0 TVP and ending with 0 flow. Maximum TVP is the maximum pressure gradient recorded across the valve from the aortic and ventricular pressure readings.</li>
			<li>The RMS forward flow rate (Qrms) provides a useful metric for quantifying the magnitude of forward flow rate as follows: 
			<img alt="Equation 1" fo:src="/files/ftp_upload/50335/50335eq1highres.jpg" src="/files/ftp_upload/50335/50335eq1.jpg" /> 
			Where &#39;n&#39; is the total number of time points collected, &#39;Qi&#39; is the instantaneous flow rate measurement collected in order &#39;i&#39;.</li>
			<li>The aortic forward, closing and leakage volumes are computed based on the following time intervals, Forward: beginning of forward flow through the valve (to), to the end of forward flow (t1); Closing: from t1 till the instance of valve closure (t2); Leakage: from t2 till the end of the cardiac cycle (t3). Total regurgitant volume is simply the sum of closing and leakage volumes.</li>
			<li>The EOA based on blood properties can be computed for the 3 intervals, P, F and H from the mean TVP during each of these periods as20: 
			<img alt="Equation 1" fo:src="/files/ftp_upload/50335/50335eq2highres.jpg" src="/files/ftp_upload/50335/50335eq2.jpg" /></li>
			<li>Energy losses are defined as follows21: 
			<img alt="Equation 1" fo:src="/files/ftp_upload/50335/50335eq3highres.jpg" src="/files/ftp_upload/50335/50335eq3.jpg" /></li>
		</ol>
		</li>
	</ol>
	</li>
</ol>

<ol>
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S., Jouven, X. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rheumatic+heart+disease.">Rheumatic heart disease.</a> Lancet. 379, 953-964 (2012).</li><li>Karaci, A. 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=Surgical+treatment+of+infective+valve+endocarditis+in+children+with+congenital+heart+disease.">Surgical treatment of infective valve endocarditis in children with congenital heart disease.</a> J. Card. Surg. 27, 93-98 (2012).</li><li>Knirsch, W., Nadal, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Infective+endocarditis+in+congenital+heart+disease.">Infective endocarditis in congenital heart disease.</a> Eur. J. Pediatr. 170, 1111-1127 (2011).</li><li>Korossis, S. 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The authors have nothing to disclose.

In this study, we have demonstrated the utility of modifying a commercially available pulsatile duplicator unit to accommodate tri-leaflet valve geometries so that hydrodynamic testing of polymer and native porcine valves can be performed. Specifically in our case, the system modified was a ViVitro left heart and systemic simulator system (Figure 1a) controlled via the ViViTest data acquisition system (ViVitro Systems, Inc, Victoria, BC, Canada). However, the system is not unlike several in vitro</em...

Heart valve disease often results from degenerative valve calcification1, rheumatic fever2, endocarditis3,4 or congenital birth defects. When valve damage occurs, causing stenosis and/or regurgitation valve prolapse and cannot be surgically repaired, the native valve is usually replaced by a prosthetic valve. Currently available options include mechanical valves (cage-ball valves, tilting disk valves, etc.), homograft, and bioprosthetic valves (porcine and bovine valves). Mechanical valves are often recommended for younger patients based on their durability; however the patient is required to remain on anticoagulant therapy to prevent thrombotic complications5. Homograft and biological prosthetic valves have been effective choices to avoid blood thinner therapy; however, these valves have elevated risk for fibrosis, calcification, degeneration, and immunogenic complications leading to valve failure6. Tissue-engineered valves are being investigated as an emerging technology7-9, but much still remains to be uncovered. Alternative durable, biocompatible, prosthetic valves are needed to improve the quality of life of the heart valve disease patients. Again, this valve design could replace the bioprosthesis used in transcatheter valve technology, with transcatheter approaches showing the potential for transforming the treatment of selected patients with heart valve disease10.
As stated by current standards, a successful heart valve substitute should have the following performance characteristics: &#34;1) allows forward flow with acceptably small mean pressure difference drop; 2) prevents retrograde flow with acceptably small regurgitation; 3) resists embolization; 4) resists hemolysis; 5) resists thrombus formation; 6) is biocompatible; 7) is compatible with in vivo diagnostic techniques; 8) is deliverable and implantable in the target population; 9) remains fixed once placed; 10) has an acceptable noise level; 11) has reproducible function; 12) maintains its functionality for a reasonable lifetime, consistent with its generic class; 13) maintains its functionality and sterility for a reasonable shelf life prior to implantation.&#34;11. Some of the shortcomings of existing valve prostheses may potentially be overcome by a polymer valve. Biocompatible polymers have been considered top candidates based on biostability, anti-hydrolysis, anti-oxidation, and advantageous mechanical properties such as high strength and viscoelasticity. In particular, elastomeric polymers may provide material deformation resembling native valve dynamics. Elastomers can be tailored to mimic soft tissue properties, and they may be the only artificial materials available that are bio-tolerant and that can withstand the coupled, in vivo, fluid-induced, flexural and tensile stresses, yet, move in a manner resembling healthy, native valve motion. Moreover, elastomers can be mass-produced in a variety of sizes, stored with ease, are expected to be cost-effective devices and can be structurally augmented with fibrous reinforcement.
The concept of the use of polymer materials to assemble a tri-leaflet valve is not new and has been the subject of several research investigations over the last 50 years12, which were abandoned largely due to limited valve durability. However, with the advent of novel manufacturing methodologies13,14, the reinforcement of polymer materials15,16 and potentially seamless integration of polymer valve substitutes with transcatheter valve technology, there has recently been a renewed interest and activity in developing polymer valves as a potentially viable alternative to currently available commercial valves. In this light, a protocol for enabling testing of these valves to assess hydrodynamic functionality is the first step in the evaluation process; yet commercially available pulse simulator systems generally do not come equipped to accommodate tri-leaflet valve designs and contain an annular spacing to insert commercially available heart valves (e.g. tilting disc, bi-leaflet mechanical heart valves). Secondly, polymer valves are an emerging technology whose hydrodynamics can only be assessed in a relative context. Even though native heart valve pressure and flow data is available, it is important to conduct testing of native aortic porcine valves, which are biologically similar to human valves, using the same pulsatile simulator that is used to evaluate the polymer valves so as to account for measurement differences that may be system dependent. Thus, the goal of this study was to demonstrate how a commercially available pulse simulator can be fitted with an assembly to accommodate tri-leaflet valve constructs and to systematically evaluate polymer valve hydrodynamic metrics in a relative context in comparison to mechanical and native porcine heart valve counterparts. In our case, novel tri-leaflet silicone polymer valves previously developed at the University of Florida 13 comprised the polymer valve group.

<table border="1">
	<tbody>
		<tr>
			<td>Name of Reagent/Material</td>
			<td>Company</td>
			<td>Catalog Number</td>
			<td>Comments</td>
		</tr>
		<tr>
			<td>Pump</td>
			<td>ViVitro Labs</td>
			<td></td>
			<td><a href="http://vivitrolabs.com/products/superpump/" target="_blank">http://vivitrolabs.com/products/superpump/</a></td>
		</tr>
		<tr>
			<td>Flow Meter and Probe</td>
			<td>Carolina Medical</td>
			<td>Model 501D</td>
			<td><a href="http://www.carolinamedicalelectronics.com/documents/FM501.pdf" target="_blank">http://www.carolinamedicalelectronics.com/documents/FM501.pdf</a></td>
		</tr>
		<tr>
			<td>Pressure Transducer</td>
			<td>ViVitro Labs</td>
			<td>HCM018</td>
			<td></td>
		</tr>
		<tr>
			<td>ViVitro Pressure Measuring Assembly</td>
			<td>ViVitro Labs</td>
			<td>6186</td>
			<td></td>
		</tr>
		<tr>
			<td>Valve holder</td>
			<td>WB Engineering</td>
			<td></td>
			<td>Designed by Florida International University. Manufactured by WB Engineering</td>
		</tr>
		<tr>
			<td>Pulse Duplicator</td>
			<td>ViVitro Labs</td>
			<td>PD2010</td>
			<td><a href="http://vivitrolabs.com/wp-content/uploads/Pulse-Duplicator-Accessories1.pdf" target="_blank">http://vivitrolabs.com/wp-content/uploads/Pulse-Duplicator-Accessories1.pdf</a></td>
		</tr>
		<tr>
			<td>Pulse Duplicator Data Acquisition and Control System, including ViViTest Software</td>
			<td>ViVitro Labs</td>
			<td>PDA2010</td>
			<td><a href="http://vivitrolabs.com/products/software-daq" target="_blank">http://vivitrolabs.com/products/software-daq</a></td>
		</tr>
		<tr>
			<td>Porcine Hearts and Native Aortic Valves</td>
			<td>Mary&#39;s Ranch Inc</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Bi-leaflet Mechanical Valves</td>
			<td>Saint Jude Medical</td>
			<td></td>
			<td><a href="http://www.sjm.com/" target="_blank">http://www.sjm.com/</a></td>
		</tr>
		<tr>
			<td>High Vacuum Grease</td>
			<td>Dow Corning Corporation</td>
			<td></td>
			<td><a href="http://www1.dowcorning.com/DataFiles/090007b281afed0e.pdf" target="_blank">http://www1.dowcorning.com/DataFiles/090007b281afed0e.pdf</a></td>
		</tr>
		<tr>
			<td>Glycerin</td>
			<td>McMaster-Carr</td>
			<td>3190K293</td>
			<td>99% Natural 5 gal</td>
		</tr>
		<tr>
			<td>Phosphate Buffered Saline (PBS)</td>
			<td>Fisher Scientific</td>
			<td>MT21031CV</td>
			<td>100 ml/heart</td>
		</tr>
		<tr>
			<td>Antimycotic/Antibiotic Solution</td>
			<td>Fisher Scientific</td>
			<td>SV3007901</td>
			<td>1 ml in 100 ml of PBS/heart; 20 ml for ViVitro System</td>
		</tr>
		<tr>
			<td>NaCl</td>
			<td>Sigma-Aldrich</td>
			<td>S3014-500G</td>
			<td>9 g/L of deionized water</td>
		</tr>
		<tr>
			<td>Deionized Water</td>
			<td>EMD Millipore Chemicals</td>
			<td></td>
			<td>Millipore Deionized Purification System. 1.3 L for ViVitro System, 200 ml for heart valve dissection process</td>
		</tr>
	</tbody>
</table>

protocol for relative hydrodynamic assessment of tri-leaflet polymer valves

Limitations of currently available prosthetic valves, xenografts, and homografts have prompted a recent resurgence of developments in the area of tri-leaflet polymer valve prostheses. However, identification of a protocol for initial assessment of polymer valve hydrodynamic functionality is paramount during the early stages of the design process. Traditional in vitro pulse duplicator systems are not configured to accommodate flexible tri-leaflet materials; in addition, assessment of polymer valve functionality needs to be made in a relative context to native and prosthetic heart valves under identical test conditions so that variability in measurements from different instruments can be avoided. Accordingly, we conducted hydrodynamic assessment of i) native (n = 4, mean diameter, D = 20 mm), ii) bi-leaflet mechanical (n= 2, D = 23 mm) and iii) polymer valves (n = 5, D = 22 mm) via the use of a commercially available pulse duplicator system (ViVitro Labs Inc, Victoria, BC) that was modified to accommodate tri-leaflet valve geometries. Tri-leaflet silicone valves developed at the University of Florida comprised the polymer valve group. A mixture in the ratio of 35:65 glycerin to water was used to mimic blood physical properties. Instantaneous flow rate was measured at the interface of the left ventricle and aortic units while pressure was recorded at the ventricular and aortic positions. Bi-leaflet and native valve data from the literature was used to validate flow and pressure readings. The following hydrodynamic metrics were reported: forward flow pressure drop, aortic root mean square forward flow rate, aortic closing, leakage&#160;and regurgitant volume, transaortic closing, leakage, and total energy losses. Representative results indicated that hydrodynamic metrics from the three valve groups could be successfully obtained by incorporating a custom-built assembly into a commercially available pulse duplicator system and subsequently, objectively compared to provide insights on functional aspects of polymer valve design.

Representative flow and pressure waveforms are shown in Figures 3, 4 and 5. The plots were averaged over the sample size of valves tested for each group, which was, n = 5, 4, and 2 valves for polymer, native porcine and bi-leaflet groups, respectively. The mean hydrodynamic metrics and the standard error of the mean for these sample sizes are presented in Table 1.
<img alt="Figure 1" src="/files/ftp_upload...

Watch this Scientific Journal Video about Protocol for Relative Hydrodynamic Assessment of Tri-leaflet Polymer Valves at JoVE.com

Protocol for Relative Hydrodynamic Assessment of Tri-leaflet Polymer Valves

There has been renewed interest in developing polymer valves. Here, the objectives are to demonstrate the feasibility of modifying a commercial pulse duplicator to accommodate tri-leaflet geometries and to define a protocol to present polymer valve hydrodynamic data in comparison to native and prosthetic valve data collected under near-identical conditions.

Limitations of currently available prosthetic valves, xenografts, and homografts have prompted a recent resurgence of developments in the area of ...

protocol-for-relative-hydrodynamic-assessment-tri-leaflet-polymer

Research

JoVE Journal

Bioengineering

13.7K visualizações.  Florida International University. Tem havido um interesse renovado no desenvolvimento de válvulas de polímeros. Aqui, os objetivos são demonstrar a viabilidade de modificação de um duplicador de pulso comercial para acomodar geometrias tri-folheto e definir um protocolo para apresentar polímero dados hidrodinâmicos de válvulas em relação aos dados de válvulas nativas e protéticas, recolhidos em condições quase idênticas.

Vídeo: Protocol for Relative Hydrodynamic Assessment of Tri-leaflet Polymer Valves

A Microfluidic-based Hydrodynamic Trap for Single Particles

The ability to confine and manipulate single particles in free solution is a key enabling technology for fundamental and applied science. Methods for particle trapping based on optical, magnetic, electrokinetic, and acoustic techniques have led to major advancements in physics and biology ranging from the molecular to cellular level. In this article, we introduce a new microfluidic-based technique for particle trapping and manipulation based solely on hydrodynamic fluid flow. Using this method, we demonstrate trapping of micro- and nano-scale particles in aqueous solutions for long time scales. The hydrodynamic trap consists of an integrated microfluidic device with a cross-slot channel geometry where two opposing laminar streams converge, thereby generating a planar extensional flow with a fluid stagnation point (zero-velocity point). In this device, particles are confined at the trap center by active control of the flow field to maintain particle position at the fluid stagnation point. In this manner, particles are effectively trapped in free solution using a feedback control algorithm implemented with a custom-built LabVIEW code. The control algorithm consists of image acquisition for a particle in the microfluidic device, followed by particle tracking, determination of particle centroid position, and active adjustment of fluid flow by regulating the pressure applied to an on-chip pneumatic valve using a pressure regulator. In this way, the on-chip dynamic metering valve functions to regulate the relative flow rates in the outlet channels, thereby enabling fine-scale control of stagnation point position and particle trapping. The microfluidic-based hydrodynamic trap exhibits several advantages as a method for particle trapping. Hydrodynamic trapping is possible for any arbitrary particle without specific requirements on the physical or chemical properties of the trapped object. In addition, hydrodynamic trapping enables confinement of a "single" target object in concentrated or crowded particle suspensions, which is difficult using alternative force field-based trapping methods. The hydrodynamic trap is user-friendly, straightforward to implement and may be added to existing microfluidic devices to facilitate trapping and long-time analysis of particles. Overall, the hydrodynamic trap is a new platform for confinement, micromanipulation, and observation of particles without surface immobilization and eliminates the need for potentially perturbative optical, magnetic, and electric fields in the free-solution trapping of small particles.

In this article, we present a microfluidic-based method for particle confinement based on hydrodynamic flow. We demonstrate stable particle trapping at a fluid stagnation point using a feedback control mechanism, thereby enabling confinement and micromanipulation of arbitrary particles in an integrated microdevice.

The ability to confine and manipulate single particles in free solution is a key enabling technology for fundamental and applied science.  Methods for ...

Design of a Cyclic Pressure Bioreactor for the Ex Vivo Study of Aortic Heart Valves

The aortic valve, located between the left ventricle and the aorta, allows for unidirectional blood flow, preventing backflow into the ventricle. Aortic valve leaflets are composed of interstitial cells suspended within an extracellular matrix (ECM) and are lined with an endothelial cell monolayer. The valve withstands a harsh, dynamic environment and is constantly exposed to shear, flexion, tension, and compression. Research has shown calcific lesions in diseased valves occur in areas of high mechanical stress as a result of endothelial disruption or interstitial matrix damage1-3. Hence, it is not surprising that epidemiological studies have shown high blood pressure to be a leading risk factor in the onset of aortic valve disease4. 
The only treatment option currently available for valve disease is surgical replacement of the diseased valve with a bioprosthetic or mechanical valve5. Improved understanding of valve biology in response to physical stresses would help elucidate the mechanisms of valve pathogenesis. In turn, this could help in the development of non-invasive therapies such as pharmaceutical intervention or prevention. Several bioreactors have been previously developed to study the mechanobiology of native or engineered heart valves6-9. Pulsatile bioreactors have also been developed to study a range of tissues including cartilage10, bone11 and bladder12. The aim of this work was to develop a cyclic pressure system that could be used to elucidate the biological response of aortic valve leaflets to increased pressure loads. 
The system consisted of an acrylic chamber in which to place samples and produce cyclic pressure, viton diaphragm solenoid valves to control the timing of the pressure cycle, and a computer to control electrical devices. The pressure was monitored using a pressure transducer, and the signal was conditioned using a load cell conditioner. A LabVIEW program regulated the pressure using an analog device to pump compressed air into the system at the appropriate rate. The system mimicked the dynamic transvalvular pressure levels associated with the aortic valve; a saw tooth wave produced a gradual increase in pressure, typical of the transvalvular pressure gradient that is present across the valve during diastole, followed by a sharp pressure drop depicting valve opening in systole. The LabVIEW program allowed users to control the magnitude and frequency of cyclic pressure. The system was able to subject tissue samples to physiological and pathological pressure conditions. This device can be used to increase our understanding of how heart valves respond to changes in the local mechanical environment.

Design of a Cyclic Pressure Bioreactor for the Ex Vivo Study of Aortic Heart Valves

A cyclic pressure bioreactor capable of subjecting heart valve tissue to physiological and pathological pressure conditions has been designed. A LabVIEW program allows users to control pressure magnitude, amplitude and frequency. This device can be used to study the mechanobiology of heart valve tissue or isolated cells.

The aortic valve, located between the left ventricle and the aorta, allows for unidirectional blood flow, preventing backflow into the ventricle. Aortic ...

A Chitosan Based, Laser Activated Thin Film Surgical Adhesive, 'SurgiLux': Preparation and Demonstration

Sutures are a 4,000 year old technology that remain the 'gold-standard' for wound closure by virtue of their repair strength (~100 KPa). However, sutures can act as a nidus for infection and in many procedures are unable to effect wound repair or interfere with functional tissue regeneration.1 Surgical glues and adhesives, such as those based on fibrin and cyanoacrylates, have been developed as alternatives to sutures for the repair of such wounds. However, current commercial adhesives also have significant disadvantages, ranging from viral and prion transfer and a lack of repair strength as with the fibrin glues, to tissue toxicity and a lack of biocompatibility for the cyanoacrylate based adhesives. Furthermore, currently available surgical adhesives tend to be gel-based and can have extended curing times which limit their application.2 Similarly, the use of UV lasers to facilitate cross-linking mechanisms in protein-based or albumin 'solders' can lead to DNA damage while laser tissue welding (LTW) predisposes thermal damage to tissues.3 Despite their disadvantages, adhesives and LTW have captured approximately 30% of the wound closure market reported to be in excess of US $5 billion per annum, a significant testament to the need for sutureless technology.4
In the pursuit of sutureless technology we have utilized chitosan as a biomaterial for the development of a flexible, thin film, laser-activated surgical adhesive termed 'SurgiLux'. This novel bioadhesive uses a unique combination of biomaterials and photonics that are FDA approved and successfully used in a variety of biomedical applications and products. SurgiLux overcomes all the disadvantages associated with sutures and current surgical adhesives (see Table 1).
In this presentation we report the relatively simple protocol for the fabrication of SurgiLux and demonstrate its laser activation and tissue weld strength. SurgiLux films adhere to collagenous tissue without chemical modification such as cross-linking and through irradiation using a comparatively low-powered (120 mW) infrared laser instead of UV light. Chitosan films have a natural but weak adhesive attraction to collagen (~3 KPa), laser activation of the chitosan based SurgiLux films emphasizes the strength of this adhesion through polymer chain interactions as a consequence of transient thermal expansion.5 Without this 'activation' process, SurgiLux films are readily removed.6-9 SurgiLux has been tested both in vitro and in vivo on a variety of tissues including nerve, intestine, dura mater and cornea. In all cases it demonstrated good biocompatibility and negligible thermal damage as a consequence of irradiation.6-10

A Chitosan Based, Laser Activated Thin Film Surgical Adhesive, &#039;SurgiLux&#039;: Preparation and Demonstration

The fabrication of a novel, flexible thin film surgical adhesive from FDA approved ingredients, chitosan and indocyanine green is described. Bonding of this adhesive to collagenous tissue through a simple activation process with a low-powered infra-red laser is demonstrated.

Sutures are a 4,000 year old technology that remain the 'gold-standard' for wound closure by virtue of their repair strength (~100 KPa). However, sutures ...

Polymer Microarrays for High Throughput Discovery of Biomaterials

The discovery of novel biomaterials that are optimized for a specific biological application is readily achieved using polymer microarrays, which allows a combinatorial library of materials to be screened in a parallel, high throughput format1. Herein is described the formation and characterization of a polymer microarray using an on-chip photopolymerization technique 2. This involves mixing monomers at varied ratios to produce a library of monomer solutions, transferring the solution to a glass slide format using a robotic printing device and curing with UV irradiation. This format is readily amenable to many biological assays, including stem cell attachment and proliferation, cell sorting and low bacterial adhesion, allowing the ready identification of 'hit' materials that fulfill a specific biological criterion3-5. Furthermore, the use of high throughput surface characterization (HTSC) allows the biological performance to be correlated with physio-chemical properties, hence elucidating the biological-material interaction6. HTSC makes use of water contact angle (WCA) measurements, atomic force microscopy (AFM), X-ray photoelectron spectroscopy (XPS) and time-of-flight secondary ion mass spectrometry (ToF-SIMS). In particular, ToF-SIMS provides a chemically rich analysis of the sample that can be used to correlate the cell response with a molecular moiety. In some cases, the biological performance can be predicted from the ToF-SIMS spectra, demonstrating the chemical dependence of a biological-material interaction, and informing the development of hit materials5,3.

A description of the formation of a polymer microarray using an on-chip photopolymerization technique. The high throughput surface characterization using atomic force microscopy, water contact angle measurements, X-ray photoelectron spectroscopy and time of flight secondary ion mass spectrometry and a cell attachment assay is also described.

The discovery of novel biomaterials that are optimized for a specific biological application is readily achieved using polymer microarrays, which allows a ...

Environmentally-controlled Microtensile Testing of Mechanically-adaptive Polymer Nanocomposites for ex vivo Characterization

Implantable microdevices are gaining significant attention for several biomedical applications1-4. Such devices have been made from a range of materials, each offering its own advantages and shortcomings5,6. Most prominently, due to the microscale device dimensions, a high modulus is required to facilitate implantation into living tissue. Conversely, the stiffness of the device should match the surrounding tissue to minimize induced local strain7-9. Therefore, we recently developed a new class of bio-inspired materials to meet these requirements by responding to environmental stimuli with a change in mechanical properties10-14. Specifically, our poly(vinyl acetate)-based nanocomposite (PVAc-NC) displays a reduction in stiffness when exposed to water and elevated temperatures (e.g. body temperature). Unfortunately, few methods exist to quantify the stiffness of materials in vivo15, and mechanical testing outside of the physiological environment often requires large samples inappropriate for implantation. Further, stimuli-responsive materials may quickly recover their initial stiffness after explantation. Therefore, we have developed a method by which the mechanical properties of implanted microsamples can be measured ex vivo, with simulated physiological conditions maintained using moisture and temperature control13,16,17. 
To this end, a custom microtensile tester was designed to accommodate microscale samples13,17 with widely-varying Young's moduli (range of 10 MPa to 5 GPa). As our interests are in the application of PVAc-NC as a biologically-adaptable neural probe substrate, a tool capable of mechanical characterization of samples at the microscale was necessary. This tool was adapted to provide humidity and temperature control, which minimized sample drying and cooling17. As a result, the mechanical characteristics of the explanted sample closely reflect those of the sample just prior to explantation.
The overall goal of this method is to quantitatively assess the in vivo mechanical properties, specifically the Young's modulus, of stimuli-responsive, mechanically-adaptive polymer-based materials. This is accomplished by first establishing the environmental conditions that will minimize a change in sample mechanical properties after explantation without contributing to a reduction in stiffness independent of that resulting from implantation. Samples are then prepared for implantation, handling, and testing (Figure 1A). Each sample is implanted into the cerebral cortex of rats, which is represented here as an explanted rat brain, for a specified duration (Figure 1B). At this point, the sample is explanted and immediately loaded into the microtensile tester, and then subjected to tensile testing (Figure 1C). Subsequent data analysis provides insight into the mechanical behavior of these innovative materials in the environment of the cerebral cortex.

Environmentally-controlled Microtensile Testing of Mechanically-adaptive Polymer Nanocomposites for ex vivo Characterization

A method is discussed by which the in vivo mechanical behavior of stimuli-responsive materials is monitored as a function of time. Samples are tested ex vivo using a microtensile tester with environmental controls to simulate the physiological environment. This work further promotes understanding the in vivo behavior of our material.

Implantable microdevices are gaining significant attention for several biomedical applications1-4. Such devices have been made from a range of materials, ...

Intra-lymph Node Injection of Biodegradable Polymer Particles

Generation of adaptive immune response relies on efficient drainage or trafficking of antigen to lymph nodes for processing and presentation of these foreign molecules to T and B lymphocytes. Lymph nodes have thus become critical targets for new vaccines and immunotherapies. A recent strategy for targeting these tissues is direct lymph node injection of soluble vaccine components, and clinical trials involving this technique have been promising. Several biomaterial strategies have also been investigated to improve lymph node targeting, for example, tuning particle size for optimal drainage of biomaterial vaccine particles. In this paper we present a new method that combines direct lymph node injection with biodegradable polymer particles that can be laden with antigen, adjuvant, or other vaccine components. In this method polymeric microparticles or nanoparticles are synthesized by a modified double emulsion protocol incorporating lipid stabilizers. Particle properties (e.g.&#160;size, cargo loading) are confirmed by laser diffraction and fluorescent microscopy, respectively. Mouse lymph nodes are then identified by peripheral injection of a nontoxic tracer dye that allows visualization of the target injection site and subsequent deposition of polymer particles in lymph nodes. This technique allows direct control over the doses and combinations of biomaterials and vaccine components delivered to lymph nodes and could be harnessed in the development of new biomaterial-based vaccines.

Lymph nodes are the immunological tissues that orchestrate immune response and are a critical target for vaccines. Biomaterials have been employed to better target lymph nodes and to control delivery of antigens or adjuvants. This paper describes a technique combining these ideas to inject biocompatible polymer particles into lymph nodes.

Generation of adaptive immune response relies on efficient drainage or trafficking of antigen to lymph nodes for processing and presentation of these ...

Manufacturing Of Robust Natural Fiber Preforms Utilizing Bacterial Cellulose as Binder

A novel method of manufacturing rigid and robust natural fiber preforms is presented here. This method is based on a papermaking process, whereby loose and short sisal fibers are dispersed into a water suspension containing bacterial cellulose. The fiber and nanocellulose suspension is then filtered (using vacuum or gravity) and the wet filter cake pressed to squeeze out any excess water, followed by a drying step. This will result in the hornification of the bacterial cellulose network, holding the loose natural fibers together.
Our method is specially suited for the manufacturing of rigid and robust preforms of hydrophilic fibers. The porous and hydrophilic nature of such fibers results in significant water uptake, drawing in the bacterial cellulose dispersed in the suspension. The bacterial cellulose will then be filtered against the surface of these fibers, forming a bacterial cellulose coating. When the loose fiber-bacterial cellulose suspension is filtered and dried, the adjacent bacterial cellulose forms a network and hornified to hold the otherwise loose fibers together.
The introduction of bacterial cellulose into the preform resulted in a significant increase of the mechanical properties of the fiber preforms. This can be attributed to the high stiffness and strength of the bacterial cellulose network. With this preform, renewable high performance hierarchical composites can also be manufactured by using conventional composite production methods, such as resin film infusion (RFI) or resin transfer molding (RTM). Here, we also describe the manufacturing of renewable hierarchical composites using double bag vacuum assisted resin infusion.

We present a novel method of manufacturing rigid and robust short natural fiber preforms using a papermaking process. Bacterial cellulose acts simultaneously as the binder for the loose fibers and provides rigidity to the fiber preforms. These preforms can be infused with a resin to produce truly green hierarchical composites.

A novel method of manufacturing rigid and robust natural fiber preforms is presented here. This method is based on a papermaking process, whereby loose ...

Culturing Mouse Cardiac Valves in the Miniature Tissue Culture System

Heart valve disease is a major burden in the Western world and no effective treatment is available. This is mainly due to a lack of knowledge of the molecular, cellular and mechanical mechanisms underlying the maintenance and/or loss of the valvular structure. 
Current models used to study valvular biology include in vitro cultures of valvular endothelial and interstitial cells. Although, in vitro culturing models provide both cellular and molecular mechanisms, the mechanisms involved in the 3D-organization of the valve remain unclear. While in vivo models have provided insight into the molecular mechanisms underlying valvular development, insight into adult valvular biology is still elusive. 
In order to be able to study the regulation of the valvular 3D-organization on tissue, cellular and molecular levels, we have developed the Miniature Tissue Culture System. In this ex vivo flow model the mitral or the aortic valve is cultured in its natural position in the heart. The natural configuration and composition of the leaflet are maintained allowing the most natural response of the valvular cells to stimuli. The valves remain viable and are responsive to changing environmental conditions. This MTCS may provide advantages on studying questions including but not limited to, how does the 3D organization affect valvular biology, what factors affect 3D organization of the valve, and which network of signaling pathways regulates the 3D organization of the valve.

Here, we present an ex vivo flow model in which murine cardiac valves can be cultured allowing the study of the biology of the valve.

Heart valve disease is a major burden in the Western world and no effective treatment is available. This is mainly due to a lack of knowledge of the ...

Synthesis of Functionalized 10-nm Polymer-coated Gold Particles for Endothelium Targeting and Drug Delivery

Gold nanoparticles (AuNPs) have been used extensively in medical research due to their size, biocompatibility, and modifiable surface. Specific targeting and drug delivery are some of the applications of these AuNPs, but endothelial extracellular matrices&#39; defensive properties hamper particle uptake. To address this issue, we describe a synthesis method for ultrasmall gold nanoparticles to improve vascular delivery, with customizable functional groups and polymer lengths for further adjustments. The protocol yields 2.5 nm AuNPs that are capped with tetrakis(hydroxymethyl)phosphonium chloride (THPC). The replacement of THPC with hetero-functional polyethylene glycol (PEG) on the surface of the AuNP increases the hydrodynamic radius to 10.5 nm while providing various functional groups on the surface. The last part of the protocol includes an optional addition of a fluorophore to allow the AuNPs to be visualized under fluorescence to track nanoparticle uptake. Dialysis and lyophilization were used to purify and isolate the AuNPs. These fluorescent nanoparticles can be visualized in both in vitro and in vivo experiments due to the biocompatible PEG coating and fluorescent probes. Additionally, the size range of these nanoparticles render them an ideal candidate for probing the glycocalyx without disrupting normal vasculature function, which may lead to improved delivery and therapeutics.

We describe a method of synthesizing biocompatible 10-nm gold nanoparticles, functionalized by coating poly-ethylene glycol onto the surface. These particles can be used in vitro and in vivo for delivering therapeutics to nanoscale cellular and extracellular spaces that are difficult to access with conventional nanoparticle sizes.

Gold nanoparticles (AuNPs) have been used extensively in medical research due to their size, biocompatibility, and modifiable surface. Specific targeting ...

Morphometric Protocol for the Objective Assessment of Blastocyst Behavior During Vitrification and Warming Steps

This article describes the noninvasive method of blastocyst morphometry based on time-lapse microphotography for the accurate monitoring of a blastocyst&#39;s volume changing during individual phases before and after vitrification. The method can be useful in searching for the most optimal timing of blastocyst exposure to different concentrations of cryoprotectants by observing blastocyst shrinkage and re-expansion in different pre- and post-vitrification phases. With this methodology, the blastocyst vitrification protocol can be optimized. For a better demonstration of the usefulness of this morphometric method, two different blastocyst preparation protocols for vitrification are compared; one with using an artificial blastocoel collapsing and one without this intervention before vitrification. Both blastocysts&#39; volume changes are followed by time-lapse microphotography and measured by photo-editing software tools. The measurements are taken every 20 seconds in previtrification phases and every 5 minutes in the post-warming period. The changes of the blastocyst dimensions per time unit are presented graphically in line diagrams. The results show a long equilibration previtrification phase in which the intact blastocyst first shrinks and then slowly refills the blastocoel, entering vitrification with a fluid-filled blastocoel. The artificially collapsed blastocyst remains in its shrunken stage through the entire equilibration phase. During the vitrification phase, it also does not change its volume. Since the blastocyst morphometry shows a constant volume of the artificially collapsed blastocysts during the previtrification step, it seems that this stage could be shorter. The described protocol provides many additional comparative parameters of blastocyst behavior during and after cryopreservation on the basis of the speed and intensity of the volume changes, the number of partial blastocoel contractions or total blastocyst collapses, and the time to a total blastocoel re-expansion or the time to hatching.

Here we present a time-lapse morphometric protocol to follow the intensity of blastocyst shrinkage and re-expansion during previtrification interventions and post-warming recovery. The application of the protocol is possible in in vitro fertilization laboratories equipped with time-lapse microscopes and is recommended in the development of an optimal blastocyst vitrification method.