The authors thank Gilbert Green for technical assistance. This work was supported by grants from NIAMS/NIH (R01-AR060484) and DOD (W81XWH-12-1-0555) awarded to S.M.Y., and from NIH (U24 CA92871 and U54 CA151838) awarded to M.G.P.

All animal studies were undertaken in compliance with the regulations of the Johns Hopkins Animal Care and Use Committee.
1. Near Infrared (NIR) Fluorescence Imaging of Skeletal Collagen Remodeling In vivo
<ol>
	<li>Photo-activation of the caged CMP and tail vein injection
	<ol>
		<li>Prepare 110 &#181;l&#160;of caged CMP solution for each mouse (20-30 g mouse weight): 100 &#181;l&#160;of solution for dosing, with an extra 10 &#181;l&#160;cushion for the possible loss during transfer and injection. Mix 11 &#181;l&#160;of IR680-Ahx-NB(GPO)9 stock solution (400 &#181;M) with 11 &#181;l&#160;of 100 &#181;M cysteine in sterile PBS solution; then bring the total volume to 110 &#181;l&#160;with 1x&#160;PBS buffer. The final peptide solution contains 4 nmol of IR680-Ahx-NB(GPO)9 and 1 nmol of cysteine in 100 &#181;l&#160;of 1x&#160;PBS solution.</li>
		<li>Slowly withdraw the peptide solution into a 0.5 ml&#160;insulin syringe with a transparent barrel and a small gauge (28-30 G) needle, and carefully remove the air bubbles. Turn on the UV lamp to allow the lamp to warm up for 2-5 min.</li>
		<li>Place the peptide-containing syringe directly under the UV lamp (365 nm, &#62;25 mW/cm2) for 5 min to allow photo-activation.</li>
		<li>Meanwhile, position the mouse in a restrainer under a heated lamp;&#160;&#160;label the mouse with a permanent marker&#160;on the tail or other identification methods such as ear tag or toe tattoo,&#160;and disinfect the tail with 70% alcohol.</li>
		<li>Immediately after the UV activation, inject 100 &#181;l&#160;of UV-activated IR680-Ahx-NB(GPO)9 solution into the tail vein, preferentially at the posterior part of the tail. Place the mouse back to the cage after injection.</li>
	</ol>
	</li>
	<li>NIR fluorescence&#160;imaging of collagen remodeling activity
	<ol>
		<li>Set the heater plate of the imaging bed of the impulse imager to 37 &#176;C using Pearl Impulse 2.0 software.</li>
		<li>Put the nude or shaved mouse into the anesthesia induction chamber containing 2% isoflurane in oxygen. Verify depth of anesthesia plane by lack of mouse movement during handling and by monitoring the frequency of respirations (~1/sec).</li>
		<li>After the mouse is anesthetized, open the Pearl imager drawer, and place the animal on the imaging bed. Quickly remove the nose cone plug, and slide the mouse&#39;s muzzle inside the nose cone to keep the mouse under anesthesia (by 2% isoflurane in oxygen delivered at 2 L/min through the nose cone) during the imaging process. Once the mouse is in place, close the drawer.</li>
		<li>When the instrument indicates &#34;ready&#34; on the Image software panel, click the &#34;700 channel,&#34; &#34;white light&#34; and &#34;85 micron&#34; boxes followed by the &#34;acquire image&#34; button. NIR (excitation 685 nm, emission 720 nm) and white light photographs will then be taken using automatic focus and exposure.</li>
		<li>When the recording is completed, open the drawer, turn the mouse, and acquire images from another angle. The mouse can be placed with its front (ventral), back (dorsal) or sides facing the camera. Narrow strips of tape can be used as restraints to stretch its limbs to get the best view of the region of interest (e.g.&#160;rib cage, ankles, and wrists).</li>
		<li>A proper time to observe skeletal uptake of IR680-CMP is between 24-96 hr post injection (h.p.i.), based on the dosage (~4 nmol/mouse). To acquire high resolution images, sacrifice the mouse by cervical dislocation while under deep anesthesia after 72 h.p.i, remove the skin (and hair) using surgical forceps and scissors, and image it by NIR fluorescence as described above.</li>
		<li>Analyze the acquired NIR fluorescence images.</li>
	</ol>
	</li>
</ol>
2. Visualization of Collagens in Ex vivo&#160;Tissue Sections
<ol>
	<li>Material preparation
	<ol>
		<li>Prepare 1 ml&#160;of 10% (w/v) BSA solution in deionized water. Thaw 1 ml&#160;of goat serum in a water bath at room temperature. Cut a few pieces of Parafilm in the size of approximately 2 cm x&#160;5 cm.</li>
		<li>Prepare 10 ml&#160;of blocking buffer by mixing 500 &#181;l&#160;of goat serum, 1 ml&#160;of 10x&#160;PBS and 8.5 ml&#160;of deionized water. Prepare 5 ml&#160;of CMP dilution buffer by diluting 50 &#181;l&#160;of 10% (w/v) BSA with 4.45 ml&#160;of deionized water and 500 &#181;l&#160;10x&#160;PBS.</li>
		<li>Dilute stock solutions of carboxyfluorescein-labeled caged CMP, CFNB(GPO)9 (480 &#181;M), to 5 &#181;M using the CMP dilution buffer. Optimal CMP concentration varies from 2.5-30 &#181;M, depending on the type of tissues and the nature of tissue samples. A volume of 100 &#181;l&#160;is recommended for staining one tissue section slide. Store the formulated CMP solutions in dark.</li>
		<li>Let the mouse cornea tissue slides equilibrate to room temperature. Incubate the slides in 1x&#160;PBS buffer for 5 min. The mouse cornea tissues used in this demonstration have been prefixed with 4% paraformaldehyde in PBS solution for 1 hr, cryopreserved in Tissue-Tek O.C.T. medium, cryosectioned to 8 &#181;m thickness, and mounted on charged glass slides.</li>
	</ol>
	</li>
	<li>Tissue staining and imaging
	<ol>
		<li>Place the slides in a humidified chamber. To each slide, apply 0.5 ml&#160;of blocking solution and incubate the slides for 30 min at room temperature. Remove the blocking solution by blotting the slides on a paper towel.</li>
		<li>Apply approximately 100 &#181;l&#160;of CFNB(GPO)9 solutions to cover the tissue sections of each slide. Incubate the slides for 2 min to allow the CMP solution to permeate the tissues.</li>
		<li>Turn on the UV lamp. When the lamp is warmed up, expose the CMP-covered tissue sections to the UV light for 6 min (365 nm, ~8 mW/cm2) to deprotect the caged CMPs. After UV exposure, cover the tissue sections of each slide with a piece of Parafilm to prevent drying. Place the slides in the humidified chamber and incubate them at 4 &#176;C for 2 hr.</li>
		<li>Prepare a 1:3,000 dilution of DAPI solution in 1x&#160;PBS. After CMP staining, gently remove the Parafilm&#160;with forceps, and blot the slides on a paper towel to remove excess CMP solution. Apply approximately 100 &#181;l&#160;of diluted DAPI solutions to each tissue slide and incubate for 1 min at room temperature.</li>
		<li>After staining, immerse the slides in 1x&#160;PBS buffer in a staining jar for 5 min. Repeat this 3x in fresh 1x&#160;PBS to wash off unbound staining agents.</li>
		<li>Add a drop of mounting medium on the tissue section and cover it with a glass cover slip while avoiding trapping air bubbles. Put the slides in a cardboard slide tray to protect them from light.</li>
		<li>Image the collagen strands (FITC channel) and cell nuclei (DAPI channel) in the tissue slides using a fluorescence microscope.</li>
	</ol>
	</li>
</ol>

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The authors declare no competing financial interests.

As can be seen in this protocol, the delivery of CMP is straightforward since the UV-activation of the caged CMPs is the only additional step to the common tail vein injection protocols19. The key is to inject the peptide probes in an uncaged and metastable single-strand state. The cage group prevents CMP from self-assembly and binding to collagens (Figure 2B) until it is removed by UV light at which point the CMP starts to fold into triple helix. The injection formulation contains cysteine, which can quickly react with the cleaved cage group during UV irradiation to minimize toxicity. Numerous mice with different strains (e.g. SKH-1 and DR-1) have been tested using this formulation, and we did not detect any behavioral or other health defects in these mice up to six months. If the injection does not follow immediately after UV exposure, the CMPs will fold into homotrimeric triple helices, which have no hybridization capacity. Figure 2C shows an extreme case where prior to injection the CMPs were fully reassembled into triple helices by long delay time (&#62;2 days). To circumvent this problem, CMP solution should be prepared in relatively low concentration (e.g. ~40 &#181;M), and injected into the mice immediately after UV-decaging. Because of the slow triple helix folding rate of the CMPs in dilute conditions (half time of refolding: &#62;50 min) and the short UV exposure and injection time (5~7 min total), most of the CMPs enter the bloodstream in single-strand form when this protocol is followed. Once injected, CMPs are expected to remain as single-strands, as the concentration is reduced by a factor of 20 in the blood pool, leading to dramatic reduction in the reassembling rate18. If injection is delayed (e.g.&#160;due to animal handling) and homotrimeric refolding is suspected, the partially assembled peptides can be reactivated by heating to above 70 &#176;C to dissociate the CMPs to single-strands, followed by prompt cooling and injection. Although less convenient, such heat-dissociated CMPs also show expected results of in vivo targeting (Figure 2D).
In this demonstration, nude mice were used for NIR fluorescence imaging because up to 50% of the NIR signal can be blocked by hair. When working with haired mouse models (especially the ones with black hairs), we recommend shaving the mouse in the region of interest prior to imaging using an electric shaver or hair remover. In Figure 1, there are false positive signals from the animal&#39;s abdominal region, which is caused by auto-fluorescence of chlorophylls in the animal diet. Such background signals can be significantly lowered by switching the mouse to purified diets approximately 4 days prior to imaging. As seen in Figures 1 and 2A, there are strong CMP uptakes in the tail spine. Therefore, to avoid artifacts resulting from imperfect tail vein injections, we recommend injecting at the posterior part of the tail so that the injection site is not captured in the fluorescence image.
The labeled CMP enables targeting and imaging of specific locations of collagen remodeling in vivo that are difficult to detect using other methods. For instance, the ELISA-based blood and urine assays are sensitive for cleaved collagen telopeptide fragments, but the method cannot localize the source of the collagen turnover, because it targets soluble antigens. The main limitations of the in vivo imaging technique using NIR labeled CMPs are depth-of-penetration attenuation and quenching by proximal pooled red cells as hemoglobin is an endogenous quencher of 680/710 nm NIR dyes. Future applications of CMP in vivo imaging include SPECT or PET imaging using CMPs labeled with direct gamma-emitting radionuclides or with positron emitters, for diagnostic of disease states involving collagen remodeling (e.g.&#160;osteoporosis, arthritis, atherosclerosis, and fibrosis). These radio-labeled CMP analogs will eliminate the limitations of signal loss due to tissue depth and presence of pooled red cells, and will allow quantitative measurements of diseased tissues. In addition, the relatively late imaging time (e.g.&#160;48-96 h.p.i.) resulting from the slow binding and clearance of CMPs could make it difficult to follow rapid changes in collagen remodeling. Such limitation could be overcome by further engineering of the binding kinetics and affinity of CMP imaging agents. Since CMP binds to denatured collagens which have little structure, the CMP-based collagen imaging is complementary to second harmonic generation microscopy which can only image collagen fibers5.
When staining tissue sections with fluorescently labeled CMPs, we found it beneficial to incubate the samples in the UV-activated peptide solutions at 4 &#176;C, because the triple helical hybridization is facilitated at lower temperature1,20. It was also found that immersing the slides in fresh PBS buffer in a staining jar is the most effective way to wash off unbound CMPs, which works much better than merely pipetting washing buffers over the samples. The CMP-collagen strand hybridization is very specific and robust; therefore the simple staining protocol presented in this video can be readily modified for costaining additional biomarkers, and for identifying degraded collagens in unfixed tissue sections. This is an effective and convenient alternative to using anti-collagen antibodies for detecting fibrous collagens in various histological samples.

Collagen, the most abundant protein in mammal, plays a critical role in tissue development and regeneration by supporting proliferation and differentiation of cells. While fibrous collagens (e.g.&#160;type I, II), in connective tissues give mechanical strength to the tissues, the network-like collagen (type IV) form basic scaffold of the basement membrane where cells attach and form organized tissues. Collagen remodeling involves both collagen degradation (by proteases) and synthesis (promoted by growth factors). Although collagen remodeling, for example in bone, is part of the normal tissue renewal process, excess remodeling activity, or its occurrence in abnormal locations, typically indicates wound healing response to an injury or chronic pathological conditions such as cancer, osteoporosis, arthritis, and fibrosis1-4. The ability to directly image collagens undergoing remodeling in vivo could lead to understanding of the progression of these diseases, as well as new diagnostics and therapeutics. For example, live imaging can provide information about the severity and the location of the diseases, and can also be used to assess the efficacy of new therapeutic agents. Multiphoton laser scanning microscopy and second harmonic generation have been applied to image fibrillar collagens for monitoring extracellular matrix remodeling in tumor in live mice5. However, this technique requires animals to be mounted with transparent dorsal skinfold chambers, which is an invasive procedure. Direct and noninvasive imaging of collagen remodeling will benefit from a probe that specifically targets collagen undergoing remodeling. Such probe is difficult to prepare since it needs to distinguish the remodeling collagens from the intact and mature collagens, which are abundant in normal tissues6.
Collagen is made up of extremely rare protein structure called triple helix, which is cleaved by proteases such as matrix metalloproteinases (MMP) during collagen remodeling. The cleaved collagen fragments lose their triple helical structure and become unfolded strands (gelatin), which are further digested by nonspecific proteases1. It was recently discovered that the collagen mimetic peptide (CMP) which has the propensity to fold into triple helical structure can specifically target collagen strands which are dissociated from its triple helical state by either heat denaturation or by enzymatic degradation1,7. The binding is primarily driven by triple-helix hybridization between monomeric CMPs and the denatured collagen strands. Because CMPs self-assemble into homotrimeric triple helices at room temperature with little driving force for collagen hybridization, a caged CMP [(GPO)4NBGPO(GPO)4, designated as NB(GPO)9, O: hydroxyproline], was developed, which contains a photo-cleavable nitrobenzyl group (NB) attached to the central glycine of the peptide. The NB cage group sterically prevents the CMP from folding into triple helix; yet, removal of the cage group by UV irradiation immediately triggers the triple helical folding and collagen hybridization1. When monomeric CMPs labeled with near infrared (NIR) fluorophores are systemically delivered to model mice, they can specifically target and allow in vivo imaging of denatured collagens in tissues undergoing normal (e.g.&#160;in bone and cartilage) and pathological (e.g.&#160;in tumors) remodeling1.
Fluorescently labeled CMPs can also be used for imaging collagens in histological tissue sections. In histological study, harvested tissues are often preserved by fixation to keep the cellular components and overall tissue morphology from deterioration. The fixing procedures, which include heat, and treatment with organic solvents, and chemical cross-linking reagents (e.g.&#160;paraformaldehyde), denature the triple helical structure of collagen8. This denaturation generates sites for CMP hybridization. It has been shown that fluorescently labeled CMPs can specifically bind to collagens in fixed tissue sections (e.g.&#160;skin, cornea, and bone) even more effectively than an anti-collagen antibody, which allowed facile identification of pathologic conditions in fibrotic liver tissues9. The CMP targets denatured collagen strands containing amino acid sequence of triple helical motifs, which is common to all types of collagens. Therefore CMP can be considered a broad-spectrum collagen staining agent. Here, we present detailed experimental procedures for i) imaging denatured collagen strands in vivo and ii) visualizing collagens in ex vivo tissue sections using fluorescently labeled caged CMPs. A NIR tag, IR680, was conjugated to the caged CMP for live imaging, while carboxyfluorescein (CF) was used in tissue staining work for its compatibility with standard fluorescence microscopes. This protocol focuses on the imaging application of CMPs as related to collagen remodeling. Methods for CMP synthesis can be found in previous reports1,7,9-15. In this video report, imaging skeletal tissues in normal mice and tissue sections of mouse cornea were chosen for demonstration purpose; however the methods presented here can be readily applied to many pathologies and biological models involving collagen remodeling (e.g.&#160;tumors, wound healing), as well as to almost any prefixed tissue sample that contains collagens.

<table border="0" cellpadding="0" cellspacing="0" width="642">
	<colgroup>
		<col />
		<col />
		<col />
		<col />
	</colgroup>
	<tbody>
		<tr height="63">
			<td height="63" style="height:63px;width:216px;">IRdye 680RD Maleimide (IR680)</td>
			<td style="width:123px;">LI-COR</td>
			<td style="width:137px;">929-71050</td>
			<td style="width:167px;">Used for CMP labeling, protocol presented elsewhere1</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">5(6)-Carboxyfluorescein</td>
			<td style="width:123px;">Sigma</td>
			<td style="width:137px;">21877-5G-F</td>
			<td style="width:167px;">Used for CMP labeling, protocol presented elsewhere12</td>
		</tr>
		<tr height="41">
			<td height="41" style="height:41px;width:216px;">Cysteine</td>
			<td style="width:123px;">Advanced Chemtech</td>
			<td style="width:137px;">YC2200</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="61">
			<td height="61" style="height:61px;width:216px;">Isoflurane</td>
			<td style="width:123px;">Butler Veterinary Supply</td>
			<td style="width:137px;">4/4/5260</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Goat Serum</td>
			<td style="width:123px;">Sigma</td>
			<td style="width:137px;">G9023-10ML</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="41">
			<td height="41" style="height:41px;width:216px;">Albumin from bovine serum (BSA)</td>
			<td style="width:123px;">Sigma</td>
			<td style="width:137px;">A9647</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="41">
			<td height="41" style="height:41px;width:216px;">DAPI</td>
			<td style="width:123px;">Roche Applied Science</td>
			<td style="width:137px;">10236276001</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="41">
			<td height="41" style="height:41px;width:216px;">Fluoroshield mounting medium</td>
			<td style="width:123px;">Sigma</td>
			<td style="width:137px;">F6182-20ML</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="22">
			<td colspan="4" height="22" style="height:22px;width:643px;">Equipment</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Syringes</td>
			<td style="width:123px;">BD</td>
			<td style="width:137px;">329461</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">UV lamp</td>
			<td style="width:123px;">McMaster-Carr</td>
			<td style="width:137px;">1447T17</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="23">
			<td height="23" style="height:23px;width:216px;">Pearl impulse imager</td>
			<td style="width:123px;">LI-COR</td>
			<td style="width:137px;">9400</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="41">
			<td height="41" style="height:41px;width:216px;">Surgical forceps and scissors</td>
			<td style="width:123px;"></td>
			<td style="width:137px;"></td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="41">
			<td height="41" style="height:41px;width:216px;">EasyDip slide staining system</td>
			<td style="width:123px;">Light Labs</td>
			<td style="width:137px;">M900-12B</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="41">
			<td height="41" style="height:41px;width:216px;">20-place cardboard microscope slide tray</td>
			<td style="width:123px;">Light Labs</td>
			<td style="width:137px;"></td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Cover glasses</td>
			<td style="width:123px;">Fisher</td>
			<td style="width:137px;">12-545-c</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="41">
			<td height="41" style="height:41px;width:216px;">Fluorescence Microscope</td>
			<td style="width:123px;">Nikon</td>
			<td style="width:137px;">Eclipse TE2000-E</td>
			<td style="width:167px;"></td>
		</tr>
	</tbody>
</table>

imaging denatured collagen strands in vivo and ex vivo via photo-triggered hybridization of caged collagen mimetic peptides

Collagen is a major structural component of the extracellular matrix that supports tissue formation and maintenance. Although collagen remodeling is an integral part of normal tissue renewal, excessive amount of remodeling activity is involved in tumors, arthritis, and many other pathological conditions. During collagen remodeling, the triple helical structure of collagen molecules is disrupted by proteases in the extracellular environment. In addition, collagens present in many histological tissue samples are partially denatured by the fixation and preservation processes. Therefore, these denatured collagen strands can serve as effective targets for biological imaging. We previously developed a caged collagen mimetic peptide (CMP) that can be photo-triggered to hybridize with denatured collagen strands by forming triple helical structure, which is unique to collagens. The overall goals of this procedure are i)&#160;to image denatured collagen strands resulting from normal remodeling activities in vivo, and ii) to visualize collagens in ex vivo tissue sections using the photo-triggered caged CMPs. To achieve effective hybridization and successful in vivo and ex vivo imaging, fluorescently labeled caged CMPs are either photo-activated immediately before intravenous injection, or are directly activated on tissue sections. Normal skeletal collagen remolding in nude mice and collagens in prefixed mouse cornea tissue sections are imaged in this procedure. The imaging method based on the CMP-collagen hybridization technology presented here could lead to deeper understanding of the tissue remodeling process, as well as allow development of new diagnostics for diseases associated with high collagen remodeling activity.

Figure 1 shows a typical result of how photo-triggered IR680-Ahx-(GPO)9 distributes in a healthy female SKH1 nude mouse after 24-96 hr post injection. It shows apparent CMP accumulation in the skeleton after 24 h.p.i., at which point most of unbound peptides have been largely cleared out. The process of CMP hybridizing with the remodeling collagen strands seems relatively slow, especially in contrast to other peptide imaging probes (e.g.&#160;RGD16). This most likely is because of the slow folding rate of the collagen triple helix17,18. However, once hybridized, the CMP strands are strongly bound to the collagenous tissues, as only slight signal reduction is seen after 24 h.p.i. (Figure 1). At 96 h.p.i., the NIR fluorescence image of the mouse after skin removal clearly demonstrates the skeletal uptake of IR680-Ahx-(GPO)9 in spine and ribs, as well as within the knees, ankles, wrists, and lower mandibles (Figure 2A).
In ex vivo tissue staining, photo-triggered fluorescently-labeled caged CMPs specifically hybridize to denatured collagen strands in tissue sections. In Figure 3, CMP staining clearly reveals the fine parallel collagen fibrils in the corneal stroma, which demonstrates its use as a collagen specific staining agent.
<img alt="Figure 1" fo:content-width="3.5in" fo:src="/files/ftp_upload/51052/51052fig1highres.jpg" src="/files/ftp_upload/51052/51052fig1.jpg" /> 
Figure 1. Serial NIR fluorescence images of a nude mouse administered intravenously with photo-decaged IR680-Ahx-(GPO)9 showing skeletal uptake over four days. <a href="https://www.jove.com/files/ftp_upload/51052/51052fig1highres.jpg" target="_blank">Click here to view larger image.</a>
<img alt="Figure 2" fo:content-width="4in" fo:src="/files/ftp_upload/51052/51052fig2highres.jpg" src="/files/ftp_upload/51052/51052fig2.jpg" /> 
Figure 2. NIR fluorescence images of mice administered with photo-decaged IR680-Ahx-(GPO)9 (A), caged IR680-Ahx-NB(GPO)9 without UV exposure (B), prefolded triple-helical [IR680-Ahx-(GPO)9]3 (C), or heat-dissociated single-strand IR680-Ahx-(GPO)9 (D) at 96 h.p.i. after skin removal. Skeletal targeting by the CMP can only be observed in A and D. NIR fluorescence signals are shown in rainbow scale. <a href="https://www.jove.com/files/ftp_upload/51052/51052fig2highres.jpg" target="_blank">Click here to view larger image.</a>
<img alt="Figure 3" fo:content-width="4in" fo:src="/files/ftp_upload/51052/51052fig3highres.jpg" src="/files/ftp_upload/51052/51052fig3.jpg" /> 
Figure 3. Fluorescence micrographs of a mouse corneal tissue section prefixed in paraformaldehyde, and stained with DAPI and photo-triggered CF(GPO)9 using Protocol 2, displaying dense collagenous stroma (stained by CMP, in green) as well as cellular epithelium and endothelium (stained by DAPI, in blue) (scale bar: 100 &#181;m).

Watch this Scientific Journal Video about Imaging Denatured Collagen Strands In vivo and Ex vivo via Photo-triggered Hybridization of Caged Collagen Mimetic Peptides at JoVE.com

Imaging Denatured Collagen Strands In vivo and Ex vivo via Photo-triggered Hybridization of Caged Collagen Mimetic Peptides

This procedure demonstrates in vivo near IR fluorescence imaging of collagen remodeling activities in mice as well as ex vivo staining of collagens in tissue sections using caged collagen mimetic peptides that can be photo-triggered to hybridize with denatured collagen strands.

Imaging Denatured Collagen Strands In vivo and Ex vivo via Photo-triggered Hybridization of Caged Collagen Mimetic Peptides

Collagen is a major structural component of the extracellular matrix that supports tissue formation and maintenance. Although collagen remodeling is an ...

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Research

JoVE Journal

Bioengineering

11.8K Views.  University of Utah. This procedure demonstrates in vivo near IR fluorescence imaging of collagen remodeling activities in mice as well as ex vivo staining of collagens in tissue sections using caged collagen mimetic peptides that can be photo-triggered to hybridize with denatured collagen strands.

Video: Imaging Denatured Collagen Strands In vivo and Ex vivo via Photo-triggered Hybridization of Caged Collagen Mimetic Peptides

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 ...

Ex vivo Mimicry of Normal and Abnormal Human Hematopoiesis

Hematopoietic stem cells require a unique microenvironment in order to sustain blood cell formation1; the bone marrow (BM) is a complex three-dimensional (3D) tissue wherein hematopoiesis is regulated by spatially organized cellular microenvironments termed niches2-4. The organization of the BM niches is critical for the function or dysfunction of normal or malignant BM5. Therefore a better understanding of the in vivo microenvironment using an ex vivo mimicry would help us elucidate the molecular, cellular and microenvironmental determinants of leukemogenesis6.
Currently, hematopoietic cells are cultured in vitro in two-dimensional (2D) tissue culture flasks/well-plates7 requiring either co-culture with allogenic or xenogenic stromal cells or addition of exogenous cytokines8. These conditions are artificial and differ from the in vivo microenvironment in that they lack the 3D cellular niches and expose the cells to abnormally high cytokine concentrations which can result in differentiation and loss of pluripotency9,10.
Herein, we present a novel 3D bone marrow culture system that simulates the in vivo 3D growth environment and supports multilineage hematopoiesis in the absence of exogenous growth factors. The highly porous scaffold used in this system made of polyurethane (PU), facilitates high-density cell growth across a higher specific surface area than the conventional monolayer culture in 2D11. Our work has indicated that this model supported the growth of human cord blood (CB) mononuclear cells (MNC)12 and primary leukemic cells in the absence of exogenous cytokines. This novel 3D mimicry provides a viable platform for the development of a human experimental model to study hematopoiesis and to explore novel treatments for leukemia.

Ex vivo Mimicry of Normal and Abnormal Human Hematopoiesis

A 3D culture system for hematopoiesis is described using human cord blood and leukemic bone marrow cells. The method is based on the use of a porous synthetic polyurethane scaffold coated with extracellular matrix proteins. This scaffold is adaptable to accommodate a wide range of cells.

Hematopoietic stem cells require a unique microenvironment in order to sustain blood cell formation1; the bone marrow (BM) is a complex three-dimensional ...

Viral Nanoparticles for In vivo Tumor Imaging

The use of nanomaterials has the potential to revolutionize materials science and medicine. Currently, a number of different nanoparticles are being investigated for applications in imaging and therapy. Viral nanoparticles (VNPs) derived from plants can be regarded as self-assembled bionanomaterials with defined sizes and shapes. Plant viruses under investigation in the Steinmetz lab include icosahedral particles formed by Cowpea mosaic virus (CPMV) and Brome mosaic virus (BMV), both of which are 30 nm in diameter. We are also developing rod-shaped and filamentous structures derived from the following plant viruses: Tobacco mosaic virus (TMV), which forms rigid rods with dimensions of 300 nm by 18 nm, and Potato virus X (PVX), which form filamentous particles 515 nm in length and 13 nm in width (the reader is referred to refs. 1 and 2 for further information on VNPs).
From a materials scientist's point of view, VNPs are attractive building blocks for several reasons: the particles are monodisperse, can be produced with ease on large scale in planta, are exceptionally stable, and biocompatible. Also, VNPs are "programmable" units, which can be specifically engineered using genetic modification or chemical bioconjugation methods 3. The structure of VNPs is known to atomic resolution, and modifications can be carried out with spatial precision at the atomic level4, a level of control that cannot be achieved using synthetic nanomaterials with current state-of-the-art technologies.
In this paper, we describe the propagation of CPMV, PVX, TMV, and BMV in Vigna ungiuculata and Nicotiana benthamiana plants. Extraction and purification protocols for each VNP are given. Methods for characterization of purified and chemically-labeled VNPs are described. In this study, we focus on chemical labeling of VNPs with fluorophores (e.g. Alexa Fluor 647) and polyethylene glycol (PEG). The dyes facilitate tracking and detection of the VNPs 5-10, and PEG reduces immunogenicity of the proteinaceous nanoparticles while enhancing their pharmacokinetics 8,11. We demonstrate tumor homing of PEGylated VNPs using a mouse xenograft tumor model. A combination of fluorescence imaging of tissues ex vivo using Maestro Imaging System, fluorescence quantification in homogenized tissues, and confocal microscopy is used to study biodistribution. VNPs are cleared via the reticuloendothelial system (RES); tumor homing is achieved passively via the enhanced permeability and retention (EPR) effect12. The VNP nanotechnology is a powerful plug-and-play technology to image and treat sites of disease in vivo. We are further developing VNPs to carry drug cargos and clinically-relevant imaging moieties, as well as tissue-specific ligands to target molecular receptors overexpressed in cancer and cardiovascular disease.

Viral Nanoparticles for In vivo Tumor Imaging

Plant viral nanoparticles (VNPs) are promising platforms for applications in biomedicine. Here, we describe the procedures for plant VNP propagation, purification, characterization, and bioconjugation. Finally, we show the application of VNPs for tumor homing and imaging using a mouse xenograft model and fluorescence imaging.

The use of nanomaterials has the potential to revolutionize materials science and medicine. Currently, a number of different nanoparticles are being ...

In vitro Synthesis of Native, Fibrous Long Spacing and Segmental Long Spacing Collagen

Collagen fibrils are present in the extracellular matrix of animal tissue to provide structural scaffolding and mechanical strength. These native collagen fibrils have a characteristic banding periodicity of ~67 nm and are formed in vivo through the hierarchical assembly of Type I collagen monomers, which are 300 nm in length and 1.4 nm in diameter. In vitro, by varying the conditions to which the monomer building blocks are exposed, unique structures ranging in length scales up to 50 microns can be constructed, including not only native type fibrils, but also fibrous long spacing and segmental long spacing collagen. Herein, we present procedures for forming the three different collagen structures from a common commercially available collagen monomer. Using the protocols that we and others have published in the past to make these three types typically lead to mixtures of structures. In particular, unbanded fibrils were commonly found when making native collagen, and native fibrils were often present when making fibrous long spacing collagen. These new procedures have the advantage of producing the desired collagen fibril type almost exclusively. The formation of the desired structures is verified by imaging using an atomic force microscope.

In vitro Synthesis of Native, Fibrous Long Spacing and Segmental Long Spacing Collagen

Simple and reproducible procedures are described for making three structurally distinct collagen assemblies from a common commercially available Type I collagen monomer. Native type, fibrous long spacing or segmental long spacing collagen can be constructed by varying the conditions to which the 300 nm long and 1.4 nm diameter monomer building block is exposed.

Collagen fibrils are present in the  extracellular matrix of animal tissue to provide structural scaffolding and  mechanical strength. These native ...

Engineering 3D Cellularized Collagen Gels for Vascular Tissue Regeneration

Synthetic materials are known to initiate clinical complications such as inflammation, stenosis, and infections when implanted as vascular substitutes. Collagen has been extensively used for a wide range of biomedical applications and is considered a valid alternative to synthetic materials due to its inherent biocompatibility (i.e., low antigenicity, inflammation, and cytotoxic responses). However, the limited mechanical properties and the related low hand-ability of collagen gels have hampered their use as scaffold materials for vascular tissue engineering. Therefore, the rationale behind this work was first to engineer cellularized collagen gels into a tubular-shaped geometry and second to enhance smooth muscle cells driven reorganization of collagen matrix to obtain tissues stiff enough to be handled.
The strategy described here is based on the direct assembling of collagen and smooth muscle cells (construct) in a 3D cylindrical geometry with the use of a molding technique. This process requires a maturation period, during which the constructs are cultured in a bioreactor under static conditions (without applied external dynamic mechanical constraints) for 1 or 2 weeks. The &#8220;static bioreactor&#8221; provides a monitored and controlled sterile environment (pH, temperature, gas exchange, nutrient supply and waste removal) to the constructs. During culture period, thickness measurements were performed to evaluate the cells-driven remodeling of the collagen matrix, and glucose consumption and lactate production rates were measured to monitor the cells metabolic activity. Finally, mechanical and viscoelastic properties were assessed for the resulting tubular constructs. To this end, specific protocols and a focused know-how (manipulation, gripping, working in hydrated environment, and so on) were developed to characterize the engineered tissues.

In this work, we present a technique for the rapid fabrication of living vascular tissues by direct culturing of collagen, smooth muscle cells and endothelial cells. In addition, a new protocol for the mechanical characterization of engineered vascular tissues is described.

Synthetic materials are known to initiate clinical complications such as inflammation, stenosis, and infections when implanted as vascular substitutes. ...

Planar Gradient Diffusion System to Investigate Chemotaxis in a 3D Collagen Matrix

The importance of cell migration can be seen through the development of human life. When cells migrate, they generate forces and transfer these forces to their surrounding area, leading to cell movement and migration. In order to understand the mechanisms that can alter and/or affect cell migration, one can study these forces. In theory, understanding the fundamental mechanisms and forces underlying cell migration holds the promise of effective approaches for treating diseases and promoting cellular transplantation. Unfortunately, modern chemotaxis chambers that have been developed are usually restricted to two dimensions (2D) and have complex diffusion gradients that make the experiment difficult to interpret. To this end, we have developed, and describe in this paper, a direct-viewing chamber for chemotaxis studies, which allows one to overcome modern chemotaxis chamber obstacles able to measure cell forces and specific concentration within the chamber in a 3D environment to study cell 3D migration. More compelling, this approach allows one to successfully model diffusion through 3D collagen matrices and calculate the coefficient of diffusion of a chemoattractant through multiple different concentrations of collagen, while keeping the system simple and user friendly for traction force microscopy (TFM) and digital volume correlation (DVC) analysis.

Cell migration is an important part of human development and life. In order to understand the mechanisms that can alter cell migration, we present a planar gradient diffusion system to investigate chemotaxis in a 3D collagen matrix, which allows one to overcome modern diffusion chamber limitations of existing assays.

The importance of cell migration can be seen through the development of human life. When cells migrate, they generate forces and transfer these forces to ...

Layer-by-layer Collagen Deposition in Microfluidic Devices for Microtissue Stabilization

Although microfluidics provides exquisite control of the cellular microenvironment, culturing cells within microfluidic devices can be challenging. 3D culture of cells in collagen type I gels helps to stabilize cell morphology and function, which is necessary for creating microfluidic tissue models in microdevices. Translating traditional 3D culture techniques for tissue culture plates to microfluidic devices is often difficult because of the limited channel dimensions. In this method, we describe a technique for modifying native type I collagen to generate polycationic and polyanionic collagen solutions that can be used with layer-by-layer deposition to create ultrathin collagen assemblies on top of cells cultured in microfluidic devices. These thin collagen layers stabilize cell morphology and function, as shown using primary hepatocytes as an example cell, allowing for the long term culture of microtissues in microfluidic devices.

The creation of functional microtissues within microfluidic devices requires the stabilization of cell phenotypes by adapting traditional cell culture techniques to the limited spatial dimensions in microdevices. Modification of collagen allows the layer-by-layer deposition of ultrathin collagen assemblies that can stabilize primary cells, such as hepatocytes, as microfluidic tissue models.

Although microfluidics provides exquisite control of the cellular microenvironment, culturing cells within microfluidic devices can be challenging. 3D ...

Preparation of 3D Collagen Gels and Microchannels for the Study of 3D Interactions In Vivo

Historically, most cellular processes have been studied in only 2 dimensions. While these studies have been informative about general cell signaling mechanisms, they neglect important cellular cues received from the structural and mechanical properties of the local microenvironment and extracellular matrix (ECM). To understand how cells interact within a physiological ECM, it is important to study them in the context of 3 dimensional assays. Cell migration, cell differentiation, and cell proliferation are only a few processes that have been shown to be impacted by local changes in the mechanical properties of a 3-dimensional ECM. Collagen I, a core fibrillar component of the ECM, is more than a simple structural element of a tissue. Under normal conditions, mechanical cues from the collagen network direct morphogenesis and maintain cellular structures. In diseased microenvironments, such as the tumor microenvironment, the collagen network is often dramatically remodeled, demonstrating altered composition, enhanced deposition and altered fiber organization. In breast cancer, the degree of fiber alignment is important, as an increase in aligned fibers perpendicular to the tumor boundary has been correlated to poorer patient prognosis1. Aligned collagen matrices result in increased dissemination of tumor cells via persistent migration2,3. The following is a simple protocol for embedding cells within a 3-dimensional, fibrillar collagen hydrogel. This protocol is readily adaptable to many platforms, and can reproducibly generate both aligned and random collagen matrices for investigation of cell migration, cell division, and other cellular processes in a tunable, 3-dimensional, physiological microenvironment.

Preparation of 3D Collagen Gels and Microchannels for the Study of 3D Interactions In Vivo

Collagen is a core component of the ECM, and provides essential cues for several cellular processes ranging from migration to differentiation and proliferation. Provided here is a protocol for embedding cells within 3D collagen hydrogels, and a more advanced technique for generating randomized or aligned collagen matrices using PDMS microchannels.

Historically, most cellular processes have been studied in only 2 dimensions.  While these studies have been informative about general cell signaling ...

Quantifying Fibrillar Collagen Organization with Curvelet Transform-Based Tools

Fibrillar collagens are prominent extracellular matrix (ECM) components, and their topology changes have been shown to be associated with the progression of a wide range of diseases including breast, ovarian, kidney, and pancreatic cancers. Freely available fiber quantification software tools are mainly focused on the calculation of fiber alignment or orientation, and they are subject to limitations such as the requirement of manual steps, inaccuracy in detection of the fiber edge in noisy background, or lack of localized feature characterization. The collagen fiber quantitation tool described in this protocol is characterized by using an optimal multiscale image representation enabled by curvelet transform (CT). This algorithmic approach allows for the removal of noise from fibrillar collagen images and the enhancement of fiber edges to provide location and orientation information directly from a fiber, rather than using the indirect pixel-wise or window-wise information obtained from other tools. This CT-based framework contains two separate, but linked, packages named &#8220;CT-FIRE&#8221; and &#8220;CurveAlign&#8221; that can quantify fiber organization on a global, region of interest (ROI), or individual fiber basis. This quantification framework has been developed for more than ten years and has now evolved into a comprehensive and user-driven collagen quantification platform. Using this platform, one can measure up to about thirty fiber features including individual fiber properties such as length, angle, width, and straightness, as well as bulk measurements such as density and alignment. Additionally, the user can measure fiber angle relative to manually or automatically segmented boundaries. This platform also provides several additional modules including ones for ROI analysis, automatic boundary creation, and post-processing. Using this platform does not require prior experience of programming or image processing, and it can handle large datasets including hundreds or thousands of images, enabling efficient quantification of collagen fiber organization for biological or biomedical applications.

Here, we present a protocol to use a curvelet transform-based, open-source MATLAB software tool for quantifying fibrillar collagen organization in the extracellular matrix of both normal and diseased tissues. This tool can be applied to images with collagen fibers or other types of line-like structures.

Fibrillar collagens are prominent extracellular matrix (ECM) components, and their topology changes have been shown to be associated with the progression ...

The Preparation of Chicken Ex Ovo Embryos and Chorioallantoic Membrane Vessels as In Vivo Model for Contrast-Enhanced Ultrasound Imaging and Microbubble-Mediated Drug Delivery Studies

The chicken embryo and the blood-vessel rich chorioallantoic membrane (CAM) is a valuable in vivo model to investigate biomedical processes, new ultrasound pulsing schemes, or novel transducers for contrast-enhanced ultrasound imaging and microbubble-mediated drug delivery. The reasons for this are the accessibility of the embryo and vessel network of the CAM as well as the low costs of the model. An important step to get access to the embryo and CAM vessels is to take the egg content out of the eggshell. In this protocol, three methods for taking the content out of the eggshell between day 5 and&#160;8 of incubation are described thus allowing the embryos to develop inside the eggshell up to these days. The described methods only require simple tools and equipment and yield a higher survival success rate of 90% for 5-day, 75% for 6-day, 50% for 7-day, and 60% for 8-day old incubated eggs in comparison to ex ovo cultured embryos (~50%). The protocol also describes how to inject cavitation nuclei, such as microbubbles, into the CAM vascular system, how to separate the membrane containing the embryo and CAM from the rest of the egg content for optically transparent studies, and how to use the chicken embryo and CAM in a variety of short-term ultrasound experiments. The in vivo&#160;chicken embryo and CAM model is extremely relevant to investigate novel imaging protocols, ultrasound contrast agents, and ultrasound pulsing schemes for contrast-enhanced ultrasound imaging, and to unravel the mechanisms of ultrasound-mediated drug delivery.

This protocol describes three methods on how to obtain and use 5 to 8-day old chicken embryos and their chorioallantoic membrane (CAM) as an in vivo&#160;model to study contrast-enhanced ultrasound imaging and microbubble-mediated drug delivery.