The authors would like to thank Mr. Sven Holder and Mr. Stephen Conley for their invaluable assistance in this study. This work was funded in part by the National Institute of Heath [Grant number R00CA134920] and the American Lung Association [Grant number RG-194681-N]. NinePoint Medical Inc. sponsored the publication costs associated with this manuscript.

1. Imaging System
The technical details of OFDI have been described previously4-6. Circumferential OFDI was conducted at imaging speeds between 25 and 100 frames per second and between 512 and 2,048 axial depth profiles per circular cross-sectional image. Custom 2.4 Fr (0.8 mm diameter) helical scanning catheters used in this study were designed to operate through the access port of standard bronchoscopes. The catheters consisted of an inner optical core to focus the light onto the bronchial wall and a single-use outer sheath. The catheter body remained stationary during imaging while the inner core was rotated at a rate between 25 and 100 Hz and translated at a pullback speed of between 1.25 and 5 mm/sec. The axial resolution of the system was 6 mm in tissue and provided an image ranging depth of 7.3 mm4-6. Catheter-based OFDI was performed in this study to replicate in vivo bronchoscopic OFDI (Figure 1). However, this protocol may also be applied to imaging with a bench-top optical system (Figure 3 and 4).
2. Imaging System Set-up
<ol>
 <li>Turn on imaging system</li>
 <li>Set and record imaging parameters (rotational speed, pullback speed, image acquisition rate, etc). For the OFDI imaging system used in this study, images were obtained at 10-50 fps.</li>
 <li>Attach catheter to rotary junction and pullback device.</li>
 <li>Spin catheter and check for image quality. Adjust system alignment and offset as needed.</li>
</ol>
3. Tissue Preparation
<ol>
 <li>Place a tabletop disposable absorbent pad on the benchtop and set lung specimen on pad.</li>
 <li>If imaging a surgical ex vivo specimen from a patient, be sure to consult the pathology department to ensure that all resection margins (bronchial, vascular, and parenchymal margins) have been assessed, documented, and/or removed by a pathologist.</li>
 <li>Identify the bronchial airway entering the resection specimen at the hilum. Remove any visible mucus within the airway using a syringe bore. If necessary, attach a longer segment of plastic tubing to the syringe bore to suction deeper within the airway.</li>
 <li>Palpate the exterior surface of the specimen to identify the lesion of interest.</li>
 <li>Using a fine metal probe, gingerly navigate through the bronchial tree until near the lesion of interest.</li>
 <li>Open the airway along the probe until the lesion of interest is visible or palpable under the airway mucosa.</li>
 <li>Carefully remove any blood or mucus from the airway mucosa overlying the lesion with a cotton-tipped applicator.</li>
 <li>Place the OFDI catheter above the airway mucosa and obtain an image to confirm the lesion is underlying the airway mucosa and to identify a high-quality imaging region of interest for histology correlation.</li>
</ol>
4. Tissue Marking
<ol>
 <li>Select the region of interest in the airway based on previous imaging findings in Step 3.8.</li>
 <li>Choose two points on the tissue along the desired line of imaging. Points may be parallel to either the longitudinal (Figure 2) or circumferential (Figure 3) aspect of the airway, depending on desired results. Space dots no more than 1.5 cm apart so that the portion of tissue may fit in one histology block for processing. If a tissue length of &gt; 1.5 cm is required, then split the tissue length into multiple 1.5 cm long inked regions of interest to create multiple matched imaging:histology pairs.</li>
 <li>Dip a fine tipped open bore needle (i.e. 25 gauge 7/8" long) into the tissue marking dye (Triangle Biomedical Sciences, Durham, NC).</li>
 <li>Carefully wipe excess ink off the outside of the needle with gauze, leaving tissue marking ink only within the needle bore.</li>
 <li>Puncture the tissue perpendicular to the airway mucosa at the chosen point along the line of imaging.</li>
 <li>Repeat steps 3.3 to 3.5 for the second point on the airway mucosa.</li>
 <li>If the ink runs over the mucosal surface away from the puncture site, use a cotton tipped applicator to carefully remove the excess ink.</li>
 <li>Remove mucus or blood on the surface of the airway mucosa with a cotton tipped applicator, if present.</li>
 <li>If the ink dots are placed circumferentially within an airway, it is useful to pin open the two sides of the airway to flatten the tissue in the imaging field (Figure 3a).</li>
</ol>
5. Imaging Tissue
<ol>
 <li>Place the OFDI catheter over each ink mark and image to ensure the marks are visible on OFDI. Marks should appear as focal disruptions within the tissue structure with overlying highly scattering particles and underlying rapid signal attenuation, which corresponds to the ink particles within the puncture site (Figure 3b, Figure 4a, Figure 4g).</li>
 <li>If ink mark(s) are not visible on OFDI, repeat steps 4.3 to 4.7 for the non-visible marks. If ink marks are visible with OFDI, proceed to step 5.3.</li>
 <li>Place the catheter parallel to the two ink marks on the airway mucosal surface such that the catheter optics overlie the tissue beyond the first ink mark (Figure 2b). Anchoring the proximal end of the catheter with a lightweight object and securing the distal end can help reduce motion artifacts.</li>
 <li>Proceed with collecting an OFDI pullback.</li>
 <li>View the OFDI pullback images to ensure both ink marks are visible in imaging and to check for motion artifacts (Figure 3 and Figure 4). If the marks are not visible, repeat steps 5.1 to 5.4.</li>
</ol>
6. Collecting and Processing Tissue
<ol>
 <li>Place a green ink dot (Triangle Biomedical Sciences, Durham, NC) on the airway mucosal tissue to orient the beginning of the imaging scan, 0.3 cm away from the ink mark that appeared first in the imaging pullback (Figure 2c).</li>
 <li>Remove tissue encompassing the two black ink marks and green ink mark. Trim tissue to fit into a standard histology processing cassette. If cutting fresh tissue is difficult, then the tissue may be fixed prior to removing the tissue for histology.</li>
 <li>Place tissue in a histology processing cassette and fix in 10% formalin for at least 48 hr.</li>
 <li>Process tissue in a tissue processor, available through any histology department.</li>
 <li>Embed tissue in paraffin such that the cut sections will be parallel to the two black ink marks on the airway surface.</li>
 <li>Use a tissue microtome to face the paraffin block until either an ink mark is visible or the entire tissue section is visible, whichever comes first.</li>
 <li>Once both black ink marks are visible, cut one 5 &mu;m thick section and mount onto a glass slide.</li>
 <li>Continue to cut and mount 5 &mu;m thick sections every 50 &mu;m until the black ink marks are no longer visible or the tissue ends, whichever comes first.</li>
 <li>Follow standard hematoxylin and eosin (H&amp;E) staining protocols to stain and coverslip slides.</li>
</ol>
7. Image Processing
If images were acquired with a benchtop scanner, or other scanning technique where both ink marks were visible in a single cross-sectional image, then the image can be directly correlated with corresponding histology. If volumetric datasets were acquired with a helical scanning catheter, the images will need to be re-interpolated so that a single 2D image bisects both ink marks for correlation with histology. This can be accomplished using ImageJ or other image processing software. In some instances, the ink may not be readily visible in which case adjacent sections/slides should be examined.

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Production and Free Access to this article is sponsored by NinePoint Medical Inc.

Assessment of early lung malignancies can be extremely challenging due to lack of symptoms and the inability to visualize early neoplastic changes radiologically or bronchoscopically. OFDI provides near histologic resolution, large area 3-dimensional views of tissue microstructure in real time2-6. Endobronchial OFDI has been demonstrated in patients as a safe technique that can be used to obtain high-resolution volumetric datasets over long airway segments in the pulmonary airways11-13 (Animation). However, only small biopsies are obtained as histopathological counterparts in the in vivo setting, which do not provide adequate correlates to OFDI for the development of imaging criteria for pulmonary pathology. In order to accurately assess OFDI features seen in pulmonary imaging, it is essential to obtain precisely matched image to histology correlations. We present a simple and effective method for precise, one to one correlation between OFDI and histology applied to airway imaging of ex vivo pulmonary resection specimens, which is applicable to nearly any ex vivo tissue type. Once imaging criteria have been established ex vivo with matched one-to-one histology, these criteria can then be applied to in vivo imaging.
The tissue dye used to mark the imaging region of interest is clearly visible in both OFDI and histology. By using straightforward techniques to orient the tissue, ink marks may be correlated in both imaging and histology to allow one to one comparisons of OFDI features and histology findings to determine the identifiable imaging characteristics of the tissue pathology. The technique is inexpensive and practical, thus making it useful in many optical imaging applications.
In the in vivo setting, methods such as laser marking may be used for tissue orientation25. However, the small size of the bronchial biopsy is still a limiting factor in using in vivo studies to develop specific imaging criteria for pulmonary pathology. Although ex vivo studies serve as an adequate alternate to in vivo imaging, there are some limitations. Ex vivo lung specimens are uninflated and often display surgically-induced atelectasis, which alters the appearance of normal alveolar structures. Inflating surgically resected lung tissue with tissue marking for histology correlation is technically challenging as most surgical lung specimens are received after pathology frozen section evaluation during which the pleural surface is disrupted, interfering with specimen inflation. Non-pathologic atelectasis is not an artifact seen in the in vivo setting, thus this limitation would not be pertinent to in vivo pulmonary imaging. Additionally, lack of blood within vessels in ex vivo specimens could make it difficult to distinguish vascular structures from other signal void structures. In the in vivo setting, the addition of Doppler OCT/OFDI26-28 to structural OCT/OFDI would aid in the identification of vessels.
Motion artifacts may be seen in vivo where they are not present ex vivo. This could be potentially problematic in standard OCT systems with slower acquisition rates. However, the rapid frame rates of OFDI systems are currently &gt; 200 fps29-31. Thus, it is not expected that motion artifact will be a significant issue. Previous in vivo OCT and OFDI imaging studies have demonstrated successful visualization of fine imaging features14,15,18,19.
In this study we have demonstrated volumetric OFDI with precise correlation to tissue-based diagnosis for evaluating pulmonary pathology. The procedure described is intended to provide precisely matched histology to be used as gold standard for the OFDI image interpretation.
Once specific imaging criteria for pulmonary pathology have been developed and validated ex vivo with matched one-to-one histology, the criteria may then be applied to subsequent in vivo imaging studies with the use of a bronchial biopsy as a gold standard assessment of the imaging features seen. This technique is presented as an application to pulmonary resection specimens, but can be applied to almost any tissue type to provide the precise imaging to histology correlation needed to determine fine imaging features of both normal and pathological tissues.

<table border="1">
 <tbody>
 <tr>
 <td>Name of the reagent</td>
 <td>Company</td>
 <td>Catalogue number</td>
 <td>Comments (optional)</td>
 </tr>
 <tr>
 <td>Tissue marking dye</td>
 <td>Triangle Biomedical</td>
 <td>TMD-BK, TMD-G</td>
 <td></td>
 </tr>
 </tbody>
</table>

optical frequency domain imaging of ex vivo pulmonary resection specimens: obtaining one to one image to histopathology correlation

Lung cancer is the leading cause of cancer-related deaths1. Squamous cell and small cell cancers typically arise in association with the conducting airways, whereas adenocarcinomas are typically more peripheral in location. Lung malignancy detection early in the disease process may be difficult due to several limitations: radiological resolution, bronchoscopic limitations in evaluating tissue underlying the airway mucosa and identifying early pathologic changes, and small sample size and/or incomplete sampling in histology biopsies. High resolution imaging modalities, such as optical frequency domain imaging (OFDI), provide non-destructive, large area 3-dimensional views of tissue microstructure to depths approaching 2 mm in real time (Figure 1)2-6. OFDI has been utilized in a variety of applications, including evaluation of coronary artery atherosclerosis6,7 and esophageal intestinal metaplasia and dysplasia6,8-10.
Bronchoscopic OCT/OFDI has been demonstrated as a safe in vivo imaging tool for evaluating the pulmonary airways11-23 (Animation). OCT has been assessed in pulmonary airways16,23 and parenchyma17,22 of animal models and in vivo human airway14,15. OCT imaging of normal airway has demonstrated visualization of airway layering and alveolar attachments, and evaluation of dysplastic lesions has been found useful in distinguishing grades of dysplasia in the bronchial mucosa11,12,20,21. OFDI imaging of bronchial mucosa has been demonstrated in a short bronchial segment (0.8 cm)18. Additionally, volumetric OFDI spanning multiple airway generations in swine and human pulmonary airways in vivo has been described19. Endobronchial OCT/OFDI is typically performed using thin, flexible catheters, which are compatible with standard bronchoscopic access ports. Additionally, OCT and OFDI needle-based probes have recently been developed, which may be used to image regions of the lung beyond the airway wall or pleural surface17.
While OCT/OFDI has been utilized and demonstrated as feasible for in vivo pulmonary imaging, no studies with precisely matched one-to-one OFDI:histology have been performed. Therefore, specific imaging criteria for various pulmonary pathologies have yet to be developed. Histopathological counterparts obtained in vivo consist of only small biopsy fragments, which are difficult to correlate with large OFDI datasets. Additionally, they do not provide the comprehensive histology needed for registration with large volume OFDI. As a result, specific imaging features of pulmonary pathology cannot be developed in the in vivo setting. Precisely matched, one-to-one OFDI and histology correlation is vital to accurately evaluate features seen in OFDI against histology as a gold standard in order to derive specific image interpretation criteria for pulmonary neoplasms and other pulmonary pathologies. Once specific imaging criteria have been developed and validated ex vivo with matched one-to-one histology, the criteria may then be applied to in vivo imaging studies. Here, we present a method for precise, one to one correlation between high resolution optical imaging and histology in ex vivo lung resection specimens. Throughout this manuscript, we describe the techniques used to match OFDI images to histology. However, this method is not specific to OFDI and can be used to obtain histology-registered images for any optical imaging technique. We performed airway centered OFDI with a specialized custom built bronchoscopic 2.4 French (0.8 mm diameter) catheter. Tissue samples were marked with tissue dye, visible in both OFDI and histology. Careful orientation procedures were used to precisely correlate imaging and histological sampling locations. The techniques outlined in this manuscript were used to conduct the first demonstration of volumetric OFDI with precise correlation to tissue-based diagnosis for evaluating pulmonary pathology24. This straightforward, effective technique may be extended to other tissue types to provide precise imaging to histology correlation needed to determine fine imaging features of both normal and diseased tissues.

The black ink marks should be between 1 - 1.5 cm apart to indicate the imaging region of interest. The green ink mark should be placed at the beginning of the imaging scan, before the first black ink mark to orient the specimen (Figure 2 and Figure 3a). Tissue ink marks should be visible on both OFDI imaging and histology (Figure 3 and 4). In normal swine (Figure 3) and human airway (Figure 4), typical airway layering should be visible. The epithelium (E) is visible as a thin, moderately signal dense, homogeneous layer at the luminal aspect of the airway. The lamina propria consists of organized signal-intense to signal-poor tissue, corresponding to various components of the lamina propria (LP) such as signal intense connective tissues including elastin and collagen (EL), and signal poor salivary-type glandular tissue (G). There are occasionally visible signal poor ducts (D) traversing the respiratory epithelium to connect with the bronchial lumen. Smooth muscle appears as discontinuous, interspersed smooth muscle fascicles and is thus not identifiable in OFDI. On the H&amp;E and trichrome stains, airway layering can be visualized (Figure 3c, 3d, 3f, 3g, 4b, 4c, 4e, and 4f), where on trichrome the superficial dense elastic and collagenous tissues appear deep blue and the underlying smooth muscle stains red (SM). Cartilage rings (C) appear as signal poor crescent-shaped structures with well-defined boundaries, which overlap in the swine airway and do not overlap in the human airway. The perichondrium surrounding the cartilage rings appears as a thin layer of signal intense tissue encompassing the signal poor cartilage rings. In the peripheral human airways (Figure 4g and 4h), alveolar attachments (A) are visible as thin, signal intense lattice-like alveolar walls with signal void alveolar spaces. Vascular spaces within the lamina propria are visible as signal void linear or circular structures with mild underlying shadowing artifact (arrows).
<img alt="Figure 1" src="/files/ftp_upload/3855/3855fig1.jpg" /> 
 Figure 1. OFDI of swine airway. In vivo images obtained from a swine airway under mechanical ventilation. (a) ODFI cross-section of proximal airway. (b) OFDI cross-section of distal airway. (c) ODFI longitudinal section of proximal airway, higher magnification image of panel e in red highlighted region. (d) OFDI longitudinal section of distal airway, higher magnification image of panel e in green highlighted region. (e) ODFI longitudinal section of airway from proximal to distal (left to right). Catheter diameter is 0.8 mm and tick marks represent 0.5 mm increments. Although different layers of the airway wall and alveolar attachments are discernible in the OFDI images, it is difficult to precisely interpret the anatomic correlate of the OFDI signals without directly registered histology. e: epithelium, lp: lamina propria, sm: submucosa, c: cartilage, a: alveolar attachments.
<img alt="Figure 2" src="/files/ftp_upload/3855/3855fig2.jpg" /> 
 Figure 2. Tissue marking of swine airway. (a) Opened airway with two black ink marks on the luminal surface placed parallel to the longitudinal aspect of the airway, 1.5 cm apart. (b) OFDI catheter placed over two black ink marks to include both marks within the OFDI pullback. (c) Airway with additional green ink mark to orient the beginning of the imaging scan on the specimen.
<img alt="Figure 3" fo:content-width="6in" fo:src="/files/ftp_upload/3855/3855fig3highres.jpg" src="/files/ftp_upload/3855/3855fig3.jpg" /> 
 Figure 3. OFDI and histology of swine airway demonstrating precise correlation using tissue marking. (a) Opened airway with two black ink marks on the luminal surface placed parallel to the circumferential aspect of the airway. Pins are used to further open the airway (arrows). (b) OFDI of swine airway with both ink marks visible (asterisks) with (c) precisely correlated histology stained with H&amp;E (asterisks: black ink marks visible on respiratory epithelium) and (d) correlated trichrome stain. Scale bar: 2 mm. (e) Higher magnification view of OFDI image with (f) corresponding histology stained with H&amp;E and (g) correlated trichrome stain. E: respiratory epithelium, EL: dense collagen and elastic tissues, SM: smooth muscle, C: cartilage rings (histological artifact has resulted in artificial separation of the cartilage rings), G: salivary gland tissue, D: salivary duct entering epithelium. Scale bar: 250 &mu;m. <a href="https://www.jove.com/files/ftp_upload/3855/3855fig3large.jpg" target="_blank"> Click here to view larger figure</a>.
<img alt="Figure 4" fo:content-width="4.75in" fo:src="/files/ftp_upload/3855/3855fig4highres.jpg" src="/files/ftp_upload/3855/3855fig4.jpg" /> 
 Figure 4. OFDI and histology of human airway demonstrating precise correlation using tissue marking. (a) OFDI of human proximal airway with both ink marks visible (asterisks). (b) Precisely correlated histology stained with H&amp;E with black ink marks visible on respiratory epithelium (asterisks) and (c) correlated trichrome stain. Scale bar: 2 mm. (d) Higher magnification view of OFDI image and (e) corresponding histology stained with H&amp;E and (f) trichrome. Scale bar: 250 &mu;m. E: respiratory epithelium, LP: lamina propria, G: salivary gland tissue, C: cartilage rings, PC: perichondrium. In the human airway, typical layering is visible. Within the loose connective tissue, there are interspersed fascicles of red-staining smooth muscle (SM, panels c and f), which do not form a continuous band and thus are not visible as a distinct layer in OFDI. (g) OFDI of human distal airway and (h) precisely correlated H&amp;E histology with black ink marks visible on respiratory epithelium (asterisks). Scale bar: 2 mm. Alveolar attachments (A) are visible as signal intense lattice-like alveolar walls with signal void alveolar spaces. Vascular spaces within the lamina propria are also visible as signal-void structures with underlying mild shadowing (arrows).

Watch this Scientific Journal Video about Optical Frequency Domain Imaging of Ex vivo Pulmonary Resection Specimens: Obtaining One to One Image to Histopathology Correlation at JoVE.com

Optical Frequency Domain Imaging of Ex vivo Pulmonary Resection Specimens: Obtaining One to One Image to Histopathology Correlation

A method to image ex vivo pulmonary resection specimens with optical frequency domain imaging (OFDI) and obtain precise correlation to histology is described, which is essential to developing specific OFDI interpretation criteria for pulmonary pathology. This method is applicable to other tissue types and imaging techniques to obtain precise imaging to histology correlation for accurate image interpretation and assessment. Imaging criteria established with this technique would then be applicable to image assessment in future in vivo studies.

Optical Frequency Domain Imaging of Ex vivo Pulmonary Resection Specimens: Obtaining One to One Image to Histopathology Correlation

Lung cancer is the leading cause of cancer-related deaths1. Squamous cell and small cell cancers typically arise in association with the conducting ...

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14.0K Views.  Harvard Medical School. A method to image ex vivo pulmonary resection specimens with optical frequency domain imaging (OFDI) and obtain precise correlation to histology is described, which is essential to developing specific OFDI interpretation criteria for pulmonary pathology. This method is applicable to other tissue types and imaging techniques to obtain precise imaging to histology correlation for accurate image interpretation and assessment. Imaging criteria established with this ...

Video: Optical Frequency Domain Imaging of Ex vivo Pulmonary Resection Specimens: Obtaining One to One Image to Histopathology Correlation

High Resolution 3D Imaging of Ex-Vivo Biological Samples by Micro CT

Non-destructive volume visualization can be achieved only by tomographic techniques, of which the most efficient is the x-ray micro computerized tomography (&mu;CT).
High resolution &mu;CT is a very versatile yet accurate (1-2 microns of resolution) technique for 3D examination of ex-vivo biological samples1, 2. As opposed to electron tomography, the &mu;CT allows the examination of up to 4 cm thick samples. This technique requires only few hours of measurement as compared to weeks in histology. In addition, &mu;CT does not rely on 2D stereologic models, thus it may complement and in some cases can even replace histological methods3, 4, which are both time consuming and destructive. Sample conditioning and positioning in &mu;CT is straightforward and does not require high vacuum or low temperatures, which may adversely affect the structure. The sample is positioned and rotated 180&deg; or 360&deg;between a microfocused x-ray source and a detector, which includes a scintillator and an accurate CCD camera, For each angle a 2D image is taken, and then the entire volume is reconstructed using one of the different available algorithms5-7. The 3D resolution increases with the decrease of the rotation step. The present video protocol shows the main steps in preparation, immobilization and positioning of the sample followed by imaging at high resolution.

High Resolution 3D Imaging of Ex-Vivo Biological Samples by Micro CT

Non-destructive volume visualization can be achieved only by tomographic techniques, of which the most efficient is the x-ray micro computerized tomography ( CT).

Non-destructive volume visualization can be achieved only by tomographic techniques, of which the most efficient is the x-ray micro computerized ...

Three-dimensional Optical-resolution Photoacoustic Microscopy

Optical microscopy, providing valuable insights at the cellular and organelle levels, has been widely recognized as an enabling biomedical technology. As the mainstays of in vivo three-dimensional (3-D) optical microscopy, single-/multi-photon fluorescence microscopy and optical coherence tomography (OCT) have demonstrated their extraordinary sensitivities to fluorescence and optical scattering contrasts, respectively. However, the optical absorption contrast of biological tissues, which encodes essential physiological/pathological information, has not yet been assessable. 
The emergence of biomedical photoacoustics has led to a new branch of optical microscopy optical-resolution photoacoustic microscopy (OR-PAM)1, where the optical irradiation is focused to the diffraction limit to achieve cellular1 or even subcellular2 level lateral resolution. As a valuable complement to existing optical microscopy technologies, OR-PAM brings in at least two novelties. First and most importantly, OR-PAM detects optical absorption contrasts with extraordinary sensitivity (i.e., 100%). Combining OR-PAM with fluorescence microscopy3 or with optical-scattering-based OCT4 (or with both) provides comprehensive optical properties of biological tissues. Second, OR-PAM encodes optical absorption into acoustic waves, in contrast to the pure optical processes in fluorescence microscopy and OCT, and provides background-free detection. The acoustic detection in OR-PAM mitigates the impacts of optical scattering on signal degradation and naturally eliminates possible interferences (i.e., crosstalks) between excitation and detection, which is a common problem in fluorescence microscopy due to the overlap between the excitation and fluorescence spectra. 
Unique for optical absorption imaging, OR-PAM has demonstrated broad biomedical applications since its invention, including, but not limited to, neurology5, 6, ophthalmology7, 8, vascular biology9, and dermatology10. In this video, we teach the system configuration and alignment of OR-PAM as well as the experimental procedures for in vivo functional microvascular imaging.

Optical-resolution photoacoustic microscopy (OR-PAM) is an emerging technology capable of imaging optical absorption contrasts in vivo with cellular resolution and sensitivity. Here, we provide a visualized instruction on the experimental protocols of OR-PAM, including system configuration, system alignment, typical in vivo experimental procedures, and functional imaging schemes.

Optical microscopy, providing valuable insights at the cellular and organelle levels, has been widely recognized as an enabling biomedical technology. As ...

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 ...

Integrated Photoacoustic Ophthalmoscopy and Spectral-domain Optical Coherence Tomography

Both the clinical diagnosis and fundamental investigation of major ocular diseases greatly benefit from various non-invasive ophthalmic imaging technologies. Existing retinal imaging modalities, such as fundus photography1, confocal scanning laser ophthalmoscopy (cSLO)2, and optical coherence tomography (OCT)3, have significant contributions in monitoring disease onsets and progressions, and developing new therapeutic strategies. However, they predominantly rely on the back-reflected photons from the retina. As a consequence, the optical absorption properties of the retina, which are usually strongly associated with retinal pathophysiology status, are inaccessible by the traditional imaging technologies.
Photoacoustic ophthalmoscopy (PAOM) is an emerging retinal imaging modality that permits the detection of the optical absorption contrasts in the eye with a high sensitivity4-7 . In PAOM nanosecond laser pulses are delivered through the pupil and scanned across the posterior eye to induce photoacoustic (PA) signals, which are detected by an unfocused ultrasonic transducer attached to the eyelid. Because of the strong optical absorption of hemoglobin and melanin, PAOM is capable of non-invasively imaging the retinal and choroidal vasculatures, and the retinal pigment epithelium (RPE) melanin at high contrasts 6,7. More importantly, based on the well-developed spectroscopic photoacoustic imaging5,8 , PAOM has the potential to map the hemoglobin oxygen saturation in retinal vessels, which can be critical in studying the physiology and pathology of several blinding diseases 9 such as diabetic retinopathy and neovascular age-related macular degeneration.
Moreover, being the only existing optical-absorption-based ophthalmic imaging modality, PAOM can be integrated with well-established clinical ophthalmic imaging techniques to achieve more comprehensive anatomic and functional evaluations of the eye based on multiple optical contrasts6,10 . In this work, we integrate PAOM and spectral-domain OCT (SD-OCT) for simultaneously in vivo retinal imaging of rat, where both optical absorption and scattering properties of the retina are revealed. The system configuration, system alignment and imaging acquisition are presented.

Photoacoustic ophthalmology (PAOM), an optical-absorption-based imaging modality, provides the complementary evaluation of the retina to the currently available ophthalmic imaging technologies. We report the using of PAOM integrated with spectral-domain optical coherence tomography (SD-OCT) for simultaneous multimodal retinal imaging in rats.

Both the clinical diagnosis and fundamental investigation of major ocular diseases greatly benefit from various non-invasive ophthalmic imaging ...

One Minute, Sub-One-Watt Photothermal Tumor Ablation Using Porphysomes, Intrinsic Multifunctional Nanovesicles

We recently developed porphysomes as intrinsically multifunctional nanovesicles. A photosensitizer, pyropheophorbide &alpha;, was conjugated to a phospholipid and then self-assembled to liposome-like spherical vesicles. Due to the extremely high density of porphyrin in the porphyrin-lipid bilayer, porphysomes generated large extinction coefficients, structure-dependent fluorescence self-quenching, and excellent photothermal efficacy. In our formulation, porphysomes were synthesized using high pressure extrusion, and displayed a mean particle size around 120 nm. Twenty-four hr post-intravenous injection of porphysomes, the local temperature of the tumor increased from 30 &deg;C to 62 &deg;C rapidly upon one minute exposure of 750 mW (1.18 W/cm2), 671 nm laser irradiation. Following the complete thermal ablation of the tumor, eschars formed and healed within 2 weeks, while in the control groups the tumors continued to grow and all reached the defined end point within 3 weeks. These data show how porphysomes can be used as potent photothermal therapy (PTT) agents.

We developed novel intrinsic multifunctional nanovesicles called porphysomes, which have structure-dependent fluorescence self-quenching and unique photothermal properties, thus functioning as potent photothermal therapy agents. We formulated porphysomes using high pressure extrusion and investigated their photothermal therapy efficacy in a xenograft tumor model.

We recently developed porphysomes as intrinsically multifunctional nanovesicles. A photosensitizer, pyropheophorbide &alpha;, was conjugated to a ...

The Use of the Ex Vivo Chandler Loop Apparatus to Assess the Biocompatibility of Modified Polymeric Blood Conduits

The foreign body reaction occurs when a synthetic surface is introduced to the body. It is characterized by adsorption of blood proteins and the subsequent attachment and activation of platelets, monocyte/macrophage adhesion, and inflammatory cell signaling events, leading to post-procedural complications. The Chandler Loop Apparatus is an experimental system that allows researchers to study the molecular and cellular interactions that occur when large volumes of blood are perfused over polymeric conduits. To that end, this apparatus has been used as an ex vivo model allowing the assessment of the anti-inflammatory properties of various polymer surface modifications. Our laboratory has shown that blood conduits, covalently modified via photoactivation chemistry with recombinant CD47, can confer biocompatibility to polymeric surfaces. Appending CD47 to polymeric surfaces could be an effective means to promote the efficacy of polymeric blood conduits. Herein is the methodology detailing the photoactivation chemistry used to append recombinant CD47 to clinically relevant polymeric blood conduits and the use of the Chandler Loop as an ex vivo experimental model to examine blood interactions with the CD47 modified and control conduits.

The Use of the Ex Vivo Chandler Loop Apparatus to Assess the Biocompatibility of Modified Polymeric Blood Conduits

Blood exposure to polymeric blood conduits initiates the foreign body reaction that has been implicated in clinical complications. Here, the Chandler Loop Apparatus, an experimental tool mimicking blood perfusion through these conduits, is described. Appendage of recombinant CD47 results in decreased evidence of the foreign body reaction on these conduits.

The foreign body reaction occurs when a synthetic surface is introduced to the body. It is characterized by adsorption of blood proteins and the ...

A Method to Estimate Cadaveric Femur Cortical Strains During Fracture Testing Using Digital Image Correlation

This protocol describes the method using digital image correlation to estimate cortical strain from high speed video images of the cadaveric femoral surface obtained from mechanical testing. This optical method requires a texture of many contrasting fiduciary marks on a solid white background for accurate tracking of surface deformation as loading is applied to the specimen. Immediately prior to testing, the surface of interest in the camera view is painted with a water-based white primer and allowed to dry for several minutes. Then, a black paint is speckled carefully over the white background with special consideration for the even size and shape of the droplets. Illumination is carefully designed and set such that there is optimal contrast of these marks while minimizing reflections through the use of filters. Images were obtained through high speed video capture at up to 12,000 frames/s. The key images prior to and including the fracture event are extracted and deformations are estimated between successive frames in carefully sized interrogation windows over a specified region of interest. These deformations are then used to compute surface strain temporally during the fracture test. The strain data is very useful for identifying fracture initiation within the femur, and for eventual validation of proximal femur fracture strength models derived from Quantitative Computed Tomography-based Finite Element Analysis (QCT/FEA).

In this protocol, the femur surface strains are estimated during fracture testing using the digital image correlation technique. The novelty of the method involves application of a high-contrast stochastic speckle pattern on the femur surface, carefully specified illumination, high speed video capture, and digital image correlation analysis for strain calculations.

This protocol describes the method using digital image correlation to estimate cortical strain from high speed video images of the cadaveric femoral ...

Rigid Embedding of Fixed and Stained, Whole, Millimeter-Scale Specimens for Section-free 3D Histology by Micro-Computed Tomography

For over a hundred years, the histological study of tissues has been the gold standard for medical diagnosis because histology allows all cell types in every tissue to be identified and characterized. Our laboratory is actively working to make technological advances in X-ray micro-computed tomography (micro-CT) that will bring the diagnostic power of histology to the study of full tissue volumes at cellular resolution (i.e., an X-ray Histo-tomography modality). Toward this end, we have made targeted improvements to the sample preparation pipeline. One key optimization, and the focus of the present work, is a straightforward method for rigid embedding of fixed and stained millimeter-scale samples. Many of the published methods for sample immobilization and correlative micro-CT imaging rely on placing the samples in paraffin wax, agarose, or liquids such as alcohol. Our approach extends this work with custom procedures and the design of a 3-dimensional printable apparatus to embed the samples in an acrylic resin directly into polyimide tubing, which is relatively transparent to X-rays. Herein, sample preparation procedures are described for the samples from 0.5 to 10 mm in diameter, which would be suitable for whole zebrafish larvae and juveniles, or other animals and tissue samples of similar dimensions. As proof of concept, we have embedded the specimens from Danio, Drosophila, Daphnia, and a mouse embryo; representative images from 3-dimensional scans for three of these samples are shown. Importantly, our methodology leads to multiple benefits including rigid immobilization, long-term preservation of laboriously-created resources, and the ability to re-interrogate samples.

We developed protocols and designed a custom apparatus to enable embedding of millimeter-scale specimens. We present sample preparation procedures with an emphasis on embedding in acrylic resin and polyimide tubing to achieve rigid immobilization and long-term storage of specimens for the interrogation of tissue architecture and cell morphology by micro-CT.

For over a hundred years, the histological study of tissues has been the gold standard for medical diagnosis because histology allows all cell types in ...

Conducting Multiple Imaging Modes with One Fluorescence Microscope

Fluorescence microscopy is a powerful tool to detect biological molecules in situ and monitor their dynamics and interactions in real-time. In addition to conventional epi-fluorescence microscopy, various imaging techniques have been developed to achieve specific experimental goals. Some of the widely used techniques include single-molecule fluorescence resonance energy transfer (smFRET), which can report conformational changes and molecular interactions with angstrom resolution, and single-molecule detection-based super-resolution (SR) imaging, which can enhance the spatial resolution approximately ten&#160;to twentyfold compared to diffraction-limited microscopy. Here we present a customer-designed integrated system, which merges multiple imaging methods in one microscope, including conventional epi-fluorescent imaging, single-molecule detection-based SR imaging, and multi-color single-molecule detection, including smFRET imaging. Different imaging methods can be achieved easily and reproducibly by switching optical elements. This set-up is easy to adopt by any research laboratory in biological sciences with a need for routine and diverse imaging experiments at a reduced cost and space relative to building separate microscopes for individual purposes.

Here we present a practical guide of building an integrated microscopy system, which merges conventional epi-fluorescent imaging, single-molecule detection-based super-resolution imaging, and multi-color single-molecule detection, including single-molecule fluorescence resonance energy transfer imaging, into one set-up in a cost-efficient way.

Fluorescence microscopy is a powerful tool to detect biological molecules in situ and monitor their dynamics and interactions in real-time. In addition to ...

Single-Cell Optical Action Potential Measurement in Human Induced Pluripotent Stem Cell-Derived Cardiomyocytes

Conventional intracellular microelectrode techniques to quantify cardiomyocyte electrophysiology are extremely complex, labor intensive, and typically carried out in low throughput. Rapid and ongoing expansion of induced pluripotent stem cell (iPSC) technology presents a new standard in cardiovascular research and alternate methods are now necessary to increase throughput of electrophysiological data at a single cell level. VF2.1Cl is a recently derived voltage sensitive dye which provides a rapid single channel, high magnitude response to fluctuations in membrane potential. It possesses kinetics superior to those of other existing voltage indicators and makes available functional data equivalent to that of traditional microelectrode techniques. Here, we demonstrate simplified, non-invasive action potential characterization in externally paced human iPSC derived cardiomyocytes using a modular and highly affordable photometry system.

Here we describe optical acquisition and characterization of action potentials from induced pluripotent stem cell derived cardiomyocytes using a high-speed modular photometry system.

Conventional intracellular microelectrode techniques to quantify cardiomyocyte electrophysiology are extremely complex, labor intensive, and typically ...