M.Z. and K.S.F. are supported by American Heart Association Scientist Development Award 17SDG33411180, and by a grant awarded under the Pitt Innovation Challenge (PInCh&#174;), through the Clinical and Translational Science Institute at the University of Pittsburgh.

The University of Pittsburgh&#39;s Institutional Animal Care and Use Committee approved all animal protocols specified in this publication prior to undertaking any of these animal experiments.
NOTE: This protocol details how to perform ex vivo biodistribution studies utilizing ex vivo optical fluorescent imaging followed by in vivo biodistribution studies by embedding the organs in paraffin, sectioning, and performing fluorescent microscopy. Although CTP is used as an example, this method can be applied to any fluorescently labeled CPP.
1. In vivo imaging
<ol>
	<li>Synthesize CTP using solid phase synthesis techniques9,10 from a peptide synthesis facility using L-amino acids with an N-terminus labeled with Cy5.5 and C-terminus amide-capped for increased stability. 
	NOTE: All synthesized CPPs should be characterized using high-performance liquid chromatography (HPLC) and/or matrix assisted laser desorption/ionization (MALDI) prior to use11.</li>
	<li>Make a 1 mM or 10 mM stock solution of CTP in dimethyl sulfoxide (DMSO), aliquot, and store at -80 &#176;C, light-protected. 
	NOTE: Peptides are delivered as lyophilized powder. Lyophilized powder can be stored long-term (6 months-2 years) in -20 &#176;C, light-protected, and under hygroscopic conditions. When aliquoted in DMSO and stored, freeze-thaw cycles should be avoided.</li>
	<li>Weigh and anesthetize 6-week-old CD1 female mice using 2 &#181;L/g of tissue weight of a ketamine/xylazine solution (100 mg/kg ketamine and 20 mg/kg xylazine) administered intramuscularly (hind leg) or intraperitoneally (lower left quadrant). 
	NOTE: Ketamine/xylazine solutions should be made fresh on the day of use. Unused ketamine/xylazine should be disposed of appropriately and volumes used versus volumes discarded recorded in a log, as ketamine is a controlled substance. Adequate level of anesthesia will be achieved in 5&#8722;7 min, as assessed by lack of response to a toe pinch.</li>
	<li>Calculate a 10 mg/kg dose of Cy5.5-CTP, dilute with phosphate buffered saline (PBS) to no more than 200 &#181;L, and inject either retro-orbitally or through tail vein injection using an insulin syringe (Figure 1A).</li>
	<li>Allow peptide to circulate for the prespecified time, ranging from 15 min-6 h.</li>
	<li>Euthanize the mouse using the high CO2 inhalational method as specified by the Institutional Animal Care and Use Committee and open the chest cavity using scissors (Figure 1B).</li>
	<li>Use scissors to place a small nick in the lateral, free wall of the right atrium and inject 3 mL of 10% buffered formalin phosphate using a 26 G needle for the dual purpose of perfusion fixing the organs of the mouse and flushing out red blood cells (Figure 1C).</li>
	<li>Dissect out heart, lung, liver, kidneys, spleen, large intestines, small intestines, bladder, ovaries/testes, and brain and place into individual wells of a 12 well plate for ex vivo optical imaging (Figure 1D).</li>
	<li>Start the image acquisition software and click the Initialize button to prepare the system for imaging samples. Open the imaging wizard, select fluorescence, then filter pair, then select the Cy5.5 dye from the list of dyes to apply 580 nm excitation and 620 nm emission filters. Select manual settings for exposure parameters then set the stage position to B, set exposure time to 1 second, set the F/stop to 8, and set the binning to small. 
	NOTE: The excitation emission filters will vary depending on the label used. The excitation and emission of the label used can be manually entered for the software to assign excitation and emission filters. Ensure that the counts are between 600-60,000 in order to avoid saturation. Depending on the system being used, automatic acquisition optimization might be available. If the system has compensation to compare images acquired with different settings, the auto settings can be used. If no compensation is available, settings will need to be determined, saved, and used consistently across samples.</li>
	<li>When the system is completely initialized, arrange the organs of interest in a 12 well plate and place it inside of the optical imaging chamber, ensuring that the samples are arranged as desired for the images. Select acquire, then select a save location for the images. Information about the mouse can be written into the edit image labels pop-up window.</li>
	<li>After imaging, remove the samples from the chamber and store in 10% buffered formalin phosphate in a volume at least 20 times the volume of the tissue in scintillation vials with light protection at room temperature (RT).</li>
	<li>Quantify the images by setting units to Radiant Efficiency then selecting the 4x3 ROI and adjusting to fit each well into one square of the ROI evenly, followed by clicking measure. Open the grid ROI measurements tab, select export, and save the measurements as a .cvs file.</li>
</ol>
2. Histology
NOTE: Follow steps 1.1&#8722;1.8 to obtain different organs. Alternatively, the same organs that were imaged can be placed in formalin immediately and used for sectioning as detailed below.
<ol>
	<li>Store each organ individually in 10% buffered formalin phosphate for a minimum of 48 h before processing for histology.</li>
	<li>When the organs are sufficiently fixed after 48 h, transfer the organs into tissue processing and embedding cassettes and place into a tissue processing machine (Figure 1E).</li>
	<li>Set the processing machine to dehydrate the tissue in 70% ethanol for 30 min, followed by 80% ethanol for 30 min, 95% ethanol for 30 min, 95% ethanol for 30 min, 100% ethanol for 15 min, 100% ethanol for 20 min, and finally 100% ethanol for 20 min.</li>
	<li>Clear the tissue in xylene 2x, with 30 min for each xylene treatment.</li>
	<li>Infiltrate the cleared tissue with paraffin wax 4x at 60 &#176;C for 30 min with each treatment.</li>
	<li>Embed in paraffin using metal molds. Fill the mold with molten paraffin (kept at 65 &#176;C), and transfer to a cold plate. As the paraffin at the bottom of the mold begins to solidify, place the organ into the paraffin (Figure 1F).</li>
	<li>Place a labeled cassette on top of the mold as a backing and overfill with molten paraffin. Allow to cool until solid. Then store the block in a -20 &#176;C freezer overnight, allowing the wax to shrink further for easy removal from the molds. 
	NOTE: The blocks can now be stored at RT with light protection.</li>
	<li>Prepare a 38 &#176;C water bath with distilled water. Set up the microtome with a blade angle of 6&#176; and a section thickness of 10 &#181;m.</li>
	<li>Mount the tissue blocks in the microtome and begin cutting until sections containing tissue are obtained. Then place the blocks face down in the water for 5 min or until the tissue has absorbed some moisture (e.g., when a thin white outline of the tissue appears in the block [Figure 1G]). 
	NOTE: The blade should be replaced frequently to ensure the quality of the sections.</li>
	<li>Place the tissue onto a flat ice block for 10 min, then return it to the microtome in the same orientation as in step 2.9. Begin taking sections and allow truncated sections to form long ribbons of 6&#8722;10 sections each. Discard suboptimal paraffin ribbons until a high-quality ribbon of sufficient length to cover a slide is produced. Optimize sectioning by adjusting cutting speeds, tissue moisturization, and water bath temperature in order to avoid shearing or wrinkling of the tissue sections. 
	NOTE: Sections that have holes where tissue fell out, rips in the tissue, or tight wrinkles and folds in the tissue should be discarded.</li>
	<li>Carefully pick the quality ribbon of choice with blunt edge forceps and float on the surface of the 38 &#176;C water bath. Let the sections sit on the surface until they smooth out, taking care to not leave them too long to prevent the paraffin from disintegrating and tearing the section apart.</li>
	<li>Float the flattened sections onto the surface of clean glass slides. Place the slides into a 65 &#176;C oven for 30 min to melt the wax. 
	NOTE: These slides can be stored at RT with light protection.</li>
	<li>Deparaffinize the slides in xylene 3x, 10 min for each treatment.</li>
	<li>Rehydrate the tissue in 100% ethanol for 5 min, followed by 95% ethanol for 5 min, 70% ethanol for 5 min, 50% ethanol for 5 min, and finally 1x tris-buffered saline (TBS) for 5 min (Figure 1H). Allow the slides to dry overnight. 
	NOTE: These slides can be stored long-term at RT as long as they are light-protected.</li>
	<li>Mount the slides with coverslips using 125 &#181;L of mounting media containing 4&#8242;,6-diamidino-2-phenylindole (DAPI), a nuclear fluorescent probe (Figure 1I). Dry slides overnight at RT, light-protected. 
	NOTE: Dried slides can be stored long-term at 4 &#176;C, light-protected.</li>
	<li>Image slides using fluorescent microscopy. 
	NOTE: Saturated images should be avoided, as they are not quantitatively useful. Adjusting the exposure and gain settings can be lowered to avoid saturation. Take care to use the same settings across the samples being compared. Avoid bleaching of the tissue by limiting exposure times.</li>
</ol>

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H., Chassaing, G., Prochiantz, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+third+helix+of+the+Antennapedia+homeodomain+translocates+through+biological+membranes.">The third helix of the Antennapedia homeodomain translocates through biological membranes.</a> Journal of Biological Chemistry. 269 (14), 10444-10450 (1994).</li><li>Schwarze, S. R., Ho, A., Vocero-Akbani, A., Dowdy, S. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+vivo+protein+transduction:+delivery+of+a+biologically+active+protein+into+the+mouse.">In vivo protein transduction: delivery of a biologically active protein into the mouse.</a> Science. 285 (5433), 1569-1572 (1999).</li><li>Zahid, M., Robbins, P. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cell-type+specific+penetrating+peptides:+therapeutic+promises+and+challenges.">Cell-type specific penetrating peptides: therapeutic promises and challenges.</a> Molecules. 20 (7), 13055-13070 (2015).</li><li>Zahid, M., Robbins, P. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Identification+and+characterization+of+tissue-specific+protein+transduction+domains+using+peptide+phage+display.">Identification and characterization of tissue-specific protein transduction domains using peptide phage display.</a> Methods in Molecular Biology. 683, 277-289 (2011).</li><li>Zahid, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Identification+of+a+cardiac+specific+protein+transduction+domain+by+in+vivo+biopanning+using+a+M13+phage+peptide+display+library+in+mice.">Identification of a cardiac specific protein transduction domain by in vivo biopanning using a M13 phage peptide display library in mice.</a> PLoS One. 5 (8), e12252 (2010).</li><li>Zahid, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cardiac+Targeting+Peptide,+a+Novel+Cardiac+Vector:+Studies+in+Bio-Distribution,+Imaging+Application,+and+Mechanism+of+Transduction.">Cardiac Targeting Peptide, a Novel Cardiac Vector: Studies in Bio-Distribution, Imaging Application, and Mechanism of Transduction.</a> Biomolecules. 8 (4), E147 (2018).</li><li>Amblard, M., Fehrentz, J. A., Martinez, J., Subra, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Methods+and+protocols+of+modern+solid+phase+Peptide+synthesis.">Methods and protocols of modern solid phase Peptide synthesis.</a> Molecular Biotechnology. 33 (3), 239-254 (2006).</li><li>Katritzky, A. R., Yoshioka, M., Narindoshvili, T., Chung, A., Johnson, J. V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fluorescent+labeling+of+peptides+on+solid+phase.">Fluorescent labeling of peptides on solid phase.</a> Organic and Biomolecular Chemistry. 6 (24), 4582-4586 (2008).</li><li>Prabhala, B. K., Mirza, O., Hojrup, P., Hansen, P. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Characterization+of+Synthetic+Peptides+by+Mass+Spectrometry.">Characterization of Synthetic Peptides by Mass Spectrometry.</a> Methods in Molecular Biology. 1348, 77-82 (2015).</li><li>Arms, L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Advantages+and+Limitations+of+Current+Techniques+for+Analyzing+the+Biodistribution+of+Nanoparticles.">Advantages and Limitations of Current Techniques for Analyzing the Biodistribution of Nanoparticles.</a> Frontiers in Pharmacology. 9, 802 (2018).</li><li>Leblond, F., Davis, S. C., Valdes, P. A., Pogue, B. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Pre-clinical+whole-body+fluorescence+imaging:+Review+of+instruments,+methods+and+applications.">Pre-clinical whole-body fluorescence imaging: Review of instruments, methods and applications.</a> Journal of Photochemistry and Photobiology. 98 (1), 77-94 (2010).</li><li>Hughes, L. D., Rawle, R. J., Boxer, S. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Choose+your+label+wisely:+water-soluble+fluorophores+often+interact+with+lipid+bilayers.">Choose your label wisely: water-soluble fluorophores often interact with lipid bilayers.</a> PLoS One. 9 (2), e87649 (2014).</li><li>McGowan, J. W., Bidwell, G. 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M.Z. and Paul D. Robbins (University of Minnesota, Minnesota, MN, USA) hold a patent on the use of cardiac targeting peptide as a cardiac vector (Cardiac-specific protein targeting domain, U.S. Patent Serial No. 9,249,184). M.Z. also serves as Chief Scientific Officer and is on the Board of Directors of the startup Vivasc Therapeutics Inc. and holds substantial future equity in it.

Animal models are essential for preclinical drug development at every stage of the process from identification of novel CPPs, biodistribution studies, mechanism of transduction, to ultimately testing for efficacy of the delivered cargo using these novel CPPs as vectors. There are many commonly used methods available to assess biodistribution such as histology, nuclear medicine imaging (SPECT and PET), and in vivo optical imaging12. Nuclear imaging methods can be cumbersome due to the limited availability of small animal SPECT and PET systems, as well as the ability to produce radiolabel-drug conjugates, which require the expertise of a radiochemist. In contrast, fluorescent labels are much simpler and can be cost effective to work with. The protocol described in this paper allows for rapid analysis of biodistribution using multiple methods. Ex vivo whole organ fluorescent optical imaging allows for the immediate comparison of fluorescence across different tissues and treatment groups and can identify organ uptake and time to peak uptake in organ of interest in a semiquantitative manner. Quantitative tissue histology requires more extensive processing to treat, section, image, and analyze tissues, but it provides more data on the microscopic level and is a current standard technique. It is worthwhile to note that direct, quantitative comparison across the two techniques is not possible, because the autofluorescence seen with each technique for different organs and excitation wavelengths varies significantly.
An important part of this protocol is selecting the right fluorophore to label candidate CPPs for experiments. One potential issue is spectra overlap, which can be problematic when multiple fluorophores are needed. The DAPI fluorescent mounting media and Cy5.5 do not have overlapping spectra. However, for certain applications where multiple fluorophores are needed, the risk of spectral overlap needs to be carefully considered. Depending on the system being used, fluorophore selection may be limited. Therefore, knowledge of the system&#39;s capabilities is key. Fluorescent optical systems are best utilized with far-red or near-infrared fluorophores due to the high tissue absorption of shorter wavelengths13. Fluorophores in the range of enhanced green fluorescent protein have a major limitation, because there is significant organ autofluorescence seen at its excitation wavelength, specifically in brain and liver tissue. Depending on the conditions of an experiment, some fluorophores are best avoided. Some water-soluble organic fluorophores have a strong interaction with lipid bilayers, which can cause false positives. Hence, taking steps to determine if a fluorophore has strong affinity to the tissue of interest is advisable14. Another factor to consider is selecting the appropriate method of fluorophore conjugation, which can be an important parameter affecting the results. CPPs can be labeled fluorescently at the N- or C-terminus through a covalent bond between the N-terminus of the peptide and the carboxyl group of the dye such as Cy5.5-NHS. Care should be taken, because the mechanism of transduction of most CPPs is not understood in detail and conjugation at one end may affect the uptake mechanism more so than at the other end. Another possibility for labeling CPPs is through biotinylating the N-terminus for conjugation to fluorescently labeled streptavidin. Using this strategy has the convenience of allowing different fluorescent streptavidin conjugates to be utilized. However, a possible limitation of this strategy is that a biotin-streptavidin complex is a large construct, which could potentially interfere with transduction.
Fluorescent optical imaging systems are an effective strategy for generating comparison of fluorescence across different organs and treatments efficiently but are incapable of producing a quantitative measure of absolute concentrations in tissue. This is due to light scattering effects within the tissue, which is further compounded by the naturally occurring variety in tissue sizes and densities, and differences in vascularity, with variable fluorescence scattering. Tissue autofluorescence can be a factor as well, due to naturally occurring biochemical sources such as collagen, or dietary sources like chlorophyll in food13.
Histology is the most commonly used method of measuring biodistribution and can potentially be used to accurately measure and compare uptake across different tissues over time. Light scattering issues are avoided using this method because all tissues are sectioned to the same thickness15. A major advantage of this method is the ability to include additional fluorescent labels postsectioning for immunohistochemistry. Although the addition of another fluorophore could make imaging more challenging, the use of fluorescent labels can be useful for localization of a CPP to particular intracellular compartments, like lysosomes or mitochondria. An antibody could be used in a confocal microscopy experiment to determine if a transduced candidate CPP colocalizes with a structure of interest, which can show the potential of a CPP as a delivery agent. One limitation of this method is that preparation of slides from organ samples can be time consuming, labor intensive, and prone to human error12,15. When imaging slides, care should be taken to not image the same location for too long to avoid photobleaching. Some photobleaching will be inevitable, depending on the sensitivity of the fluorophore. Care should be taken at every step of this protocol to protect the samples from ambient light and store them properly16. We recommend that slides be stored at 4 &#176;C, light-protected, for future imaging.
There are a variety of methods available for measuring the biodistribution of a candidate CPP that require specialized equipment and can produce comparable results, though they may require more complex CPP labeling. The protocol described in this paper uses two compatible methods to efficiently produce biodistribution data in the context of a living system while allowing for the acquisition of greater in-depth information about peptide internalization within cells from the same sample, thus cutting the number of animals needed for a study by half. These methods were used to generate the above data, which demonstrate that both methods can be utilized sequentially in the same animal, and the quality of the data generated by each. Our results also highlight the inability to directly correlate results between the two techniques in a quantitative manner.

The cell plasma membrane is a semipermeable barrier that is essential for cell integrity and survival and serves to regulate the interior of the cell by controlling movement of substances into the cell. Although vital for survival, it also presents a barrier to delivery of cargo to the cell. In 1988, the trans-activator of transcription (Tat) protein of the human immunodeficiency virus was shown to enter cultured cells and promote viral gene expression1,2, with the domains responsible for this transduction limited to an arginine and lysine rich 13-amino acid region of the third helix3 These were thus named cell penetrating peptides (CPPs). This was followed by research showing the ability of the Tat peptide to carry functional &#946;-galactosidase into multiple cell types4. Since the initial description, the number of cell-penetrating peptides has increased dramatically. These transduction domains are naturally occurring or synthetic short peptides, typically 6&#8722;20 amino acids long, that are able to carry functional cargoes across cell membranes. These cargoes can range from other small peptides, full-length proteins, nucleic acids, nanoparticles, viral particles, fluorescent molecules, and radioisotopes5. The initial CPPs described were not cell specific, with Tat being taken up by multiple cell types and even crossing the blood-brain barrier4, hence limiting its therapeutic potential. In order to identify cell-specific CPPs, investigators have undertaken phage display utilizing large, commercially available phage libraries6. Our own work using a combinatorial in vitro and in vivo phage display methodology led to the identification of a CPP named cardiac targeting peptide (CTP)7, a 12-amino acid, nonnaturally occurring peptide (NH2-APWHLSSQYSRT-COOH) that targets the heart with peak uptake at 15 min after a peripheral intravenous injection8. Using immunofluorescent colocalization with actin, an intracellular marker, and exclusion of laminin, a cell membrane marker, we showed that CTP is internalized into mouse cardiomyocytes after an intravenous injection7. Additionally, we incubated human induced pluripotent stem cell-derived beating cardiomyocytes with dual-labeled CTP, labeled with 6-carboxyfluorescein at its C-terminus and rhodamine at its N-terminus through an ester linkage that could only be cleaved off by intracellular esterases. Rapid accumulation of rhodamine into beating cardiomyocytes was observed within 15 min on confocal microscopy8.
In this article, we present two complimentary methodologies that can be used to track biodistribution and confirm tissue-specific internalization of CPPs using CTP as an example. For these methodologies, CTP was synthesized, fluorescently labeled at the N-terminus with cyanine5.5 (CY5.5), and amide-capped at the C-terminus for greater peptide stability. The two methodologies used are in vivo fluorescent optical imaging systems and fluorescent microscopy of tissue sections. Both methods are very useful in studying biodistribution, uptake, and elimination of fluorescently labeled CPPs. The advantage of assessing biodistribution using these methods over others, such as single-photon emission tomography (SPECT) or positron emission tomography (PET), is that there is no need for the time-intensive radiolabeling of CPPs compared with fluorescent labeling, which is relatively easy and in routine use at all peptide synthesis facilities. The use of in vivo imaging quickly produces biodistribution data in the context of a living system, while sectioning provides greater in-depth information about peptide uptake and localization within cells via the preservation of morphological detail. This protocol can be applied to a wide variety of organs and tissues such as the heart, lung, liver, kidney, brain, spleen, stomach, large intestine, small intestine, skeletal muscle, bone, testes/ovaries, and eyes.

&#160; &#160;

<table><tbody><tr><td>10% Buffered Formalin Phosphate</td><td>ThermoFisher</td><td>SF100-4</td><td></td></tr><tr><td>10x Tris Buffered Saline (TBS)</td><td>ThermoFisher</td><td>BP2471-1</td><td></td></tr><tr><td>12-well Cell Culture Plate</td><td>ThermoFisher</td><td>353043</td><td></td></tr><tr><td>1x TBS Solution</td><td></td><td></td><td>10x TBS is diluted 1:10 with deionized water. This solution can be stored at room temperature.</td></tr><tr><td>20 mL Scintillation Vials</td><td>Wheaton</td><td>334173</td><td></td></tr><tr><td>26G Needles</td><td>Becton Dickinson</td><td>305110</td><td></td></tr><tr><td>28G 0.5ml Insulin syringes</td><td>Becton Dickinson</td><td>329461</td><td></td></tr><tr><td>3mL Syringe</td><td>Becton Dickinson</td><td>309657</td><td></td></tr><tr><td>CD1 Mice</td><td>Charles River</td><td>022</td><td>6 to 12 week old albino, female mice</td></tr><tr><td>Cover Glass</td><td>ThermoFisher</td><td>12-544-14</td><td></td></tr><tr><td>Cy5.5-CTP-amide</td><td></td><td></td><td>Prepared by peptide synthesis, conjugated with Cy5.5 fluorophore, and purified using HPLC. Lyophilized powder is reconstituted in DMSO at 10mM concentration. After reconstituting, store at -80 &amp;deg;C.</td></tr><tr><td>Dapi Fluoromount G</td><td>SouthernBiotech</td><td>0100-20</td><td></td></tr><tr><td>Dumont #5 Forceps</td><td>Fine Science Tools</td><td>99150-20</td><td></td></tr><tr><td>Ethanol</td><td></td><td></td><td></td></tr><tr><td>Extra Fine Bonn Scissors</td><td>Fine Science Tools</td><td>14084-08</td><td></td></tr><tr><td>Ketamine HCl 100mg/mL</td><td>KetaVed</td><td>NDC 50989-161-06</td><td></td></tr><tr><td>Ketamine/Xylazine Solution</td><td></td><td></td><td>Ketamine HCl is mixed with Xylazine 1:1 to produce a stock solution containing 50mg/mL ketamine and 10mg/mL Xylazine. This solution is made fresh before each use.</td></tr><tr><td>Leica RM2235 Rotary Microtome</td><td>Leica</td><td>RM2235</td><td></td></tr><tr><td>Microscope Slides</td><td>ThermoFisher</td><td>B9992000TN</td><td></td></tr><tr><td>Paraplast X-TRA</td><td>Sigma</td><td>P3808-1KG</td><td></td></tr><tr><td>Perkin Elmer Lumina S5 IVIS</td><td>Perkin Elmer</td><td>CLS148588</td><td></td></tr><tr><td>Sparkle Optical Lens Cleaner</td><td>ThermoFisher</td><td>NC0090079</td><td></td></tr><tr><td>Tissue-Tek Processing/Embedding Cassettes</td><td>ThermoFisher</td><td>NC9499605</td><td></td></tr><tr><td>Tissue-Tek VIP Processing machine</td><td>Tissue-Tek</td><td>VIP 5A-F1</td><td></td></tr><tr><td>Xylazine 20 mg/mL</td><td>AnaSed</td><td>NDC 59399-110-20</td><td></td></tr><tr><td>Xylenes</td><td>ThermoFisher</td><td>X5-4</td><td></td></tr></tbody></table>

in vivo imaging of transduction efficiencies of cardiac targeting peptide

Since the initial description of protein transduction domains, also known as cell penetrating peptides, over 25 years ago, there has been intense interest in developing these peptides, especially cell-specific ones, as novel vectors for delivering diagnostic and therapeutic materials. Our past work involving phage display identified a novel, nonnaturally occurring, 12 amino acid-long peptide that we named cardiac targeting peptide (CTP) due to its ability to transduce normal heart tissue in vivo with peak uptake seen in as little as 15 min after an intravenous injection. We have undertaken detailed biodistribution studies by injecting CTP labeled with fluorophore cyanine5.5, allowing it to circulate for various periods of time, and euthanizing, fixing, and sectioning multiple organs followed by fluorescent microscopy imaging. In this publication, we describe these processes as well as ex vivo imaging of harvested organs using an in vivo imaging system in detail. We provide detailed methodologies and practices for undertaking transduction as well as biodistribution studies using CTP as an example.

Using this protocol, we treated three mice with a 10 mg/kg dose of Cy5.5-CTP through a retro-orbital injection (Figure 1A). After allowing the peptide to circulate for 15 min, the mice were euthanized using a CO2 chamber, the chest opened through a median sternotomy incision, the right atrium was nicked, and the mice perfusion fixed using 3 mL of 10% phosphate-buffered formalin (Figure 1B,C). After fixation, the heart, lungs, liver, kidney, spleen, and brain were dissected out and arranged in a 12 well plate for ex vivo optical fluorescent imaging (Figure 1D and Figure 2A). Three control mice with no injections were also perfused, dissected, and imaged (Figure 1B&#8722;D and Figure 2B). The fluorescence data acquired for each set of organs can be compared due to the counts being converted to radiant efficiency, the relative fluorescence unit of the imaging software. This data can be quantified to yield both qualitative images and quantitative data for each organ in question (Figure 2C) across different time points for biodistribution studies. The autofluorescence of different organs in response to different excitation wavelengths is demonstrated in Figure 3A&#8722;E. Shorter excitation wavelengths, such as enhanced green fluorescent protein (EGFP), are associated with higher autofluorescence, especially in liver and brain, than far-red (Cy5.5) or near infrared (Cy7) excitation wavelengths (Figure 3). The imaged organs were fixed in 10% phosphate buffered formalin at RT for a minimum of 48 h, followed by a transfer to tissue processing and embedding cassettes (Figure 1E). The organs were processed and paraffinized in a tissue processing machine, positioned in molds filled with paraffin, frozen on a -20 &#176;C stage, and stored overnight at -20 &#176;C (Figure 1F). The samples were sectioned (Figure 1G) and treated with solution exchanges to rehydrate the tissue (Figure 1H). The prepared slides were then mounted with DAPI fluorescent mounting medium (Figure 1I). Once dry, usually overnight, the slides were imaged using fluorescent microscopy. Representative images of each organ from a mouse are shown in Figure 4A. The images from different organs were acquired using identical settings to allow for comparison across mice, and to allow for quantification (Figure 4B).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/60895/60895fig1.jpg" /> 
Figure 1: Harvesting mouse organs for ex vivo optical fluorescent imaging and sectioning. (A) Wild type mouse injected retro-orbitally with CTP-Cy5.5. (B) Mouse dissected with chest opened, right atrium snipped, for perfusion fixation. (C) Heart injected with 3 mL of 10% buffered formalin phosphate for perfusion fixation of the animal. (D) Organs of interest harvested and arranged in a 12 well plate for fluorescent optical imaging. (E) Heart loaded into a cassette and processed using a tissue processor. (F) Heart embedded in paraffin. (G) Heart sectioned on a microtome. (H) Slides deparaffinized through a series of solution exchanges. (I) Sections mounted with coverslips using DAPI. <a href="https://www.jove.com/files/ftp_upload/60895/60895fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/60895/60895fig2v2.jpg" /> 
Figure 2: Representative images from treated and control organs. (A) Organs from a mouse treated with 10 mg/kg CTP-Cy5.5. (B) Organs from an untreated control mouse. (C) Quantification of fluorescence for each set of organs. <a href="https://www.jove.com/files/ftp_upload/60895/60895fig2v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/60895/60895fig3v2.jpg" /> 
Figure 3: Untreated mouse organs imaged at different excitation emission wavelengths to demonstrate how longer wavelengths produce less autofluorescence. (A) Cy7: 740&#8722;790. (B) Cy5.5: 660&#8722;710. (C) Cy5: 620&#8722;670. (D) Cy3: 520&#8722;570. (E) EGFP: 480&#8722;520. <a href="https://www.jove.com/files/ftp_upload/60895/60895fig3v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/60895/60895fig4.jpg" /> 
Figure 4: Transduction of heart, lung, liver, and kidney after intravenous injection in mice. Wild type mice were injected with 10 mg/kg of Cy5.5-CTP, or a random peptide (RAN-Cy5.5), and euthanized at the indicated time points. (A) Peak transduction of heart tissue was seen at 15 min with steady decrease in fluorescence over time. Some capillary uptake was noted in the lungs with robust transduction in the liver as well at the kidney glomerular capillaries, the latter implying a renal mechanism of excretion. (B) Quantification of fluorescent intensity shows significantly increased heart uptake of CTP-Cy5.5 over RAN-Cy5.5. Scale bar = 500 &#181;m. This figure has been modified from Zahid et al.8. <a href="https://www.jove.com/files/ftp_upload/60895/60895fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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In Vivo Imaging of Transduction Efficiencies of Cardiac Targeting Peptide

We describe protocols for assessing the degree of transduction by cell-penetrating peptides utilizing ex vivo imaging systems followed by paraffin embedding, sectioning, and confocal fluorescent microscopy using cardiac targeting peptide as an example. In our protocol, a single animal can be used to acquire both types of imaging assessment of the same organs, thereby cutting the number of animals needed for studies by half.

Since the initial description of protein transduction domains, also known as cell penetrating peptides, over 25 years ago, there has been intense interest ...

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Biology

5.3K Views.  University of Pittsburgh School of Medicine. We describe protocols for assessing the degree of transduction by cell-penetrating peptides utilizing ex vivo imaging systems followed by paraffin embedding, sectioning, and confocal fluorescent microscopy using cardiac targeting peptide as an example. In our protocol, a single animal can be used to acquire both types of imaging assessment of the same organs, thereby cutting the number of animals needed for studies by half.

Video: In Vivo Imaging of Transduction Efficiencies of Cardiac Targeting Peptide

Delivery of Modified mRNA in a Myocardial Infarction Mouse Model

Myocardial infarction (MI) is a leading cause of morbidity and mortality in the Western world. In the past decade, gene therapy has become a promising treatment option for heart disease, owing to its efficiency and exceptional therapeutic effects. In an effort to repair the damaged tissue post-MI, various studies have employed DNA-based or viral gene therapy but have faced considerable hurdles due to the poor and uncontrolled expression of the delivered genes, edema, arrhythmia, and cardiac hypertrophy. Synthetic modified mRNA (modRNA) presents a novel gene therapy approach that offers high, transient, safe, nonimmunogenic, and controlled mRNA delivery to the heart tissue without any risk of genomic integration. Due to these remarkable characteristics combined with its bell-shaped pharmacokinetics in the heart, modRNA has become an attractive approach for the treatment of heart disease. However, to increase its effectiveness in vivo, a consistent and reliable delivery method needs to be followed. Hence, to maximize modRNA delivery efficiency and yield consistency in modRNA use for in vivo applications, an optimized method of preparation and delivery of modRNA intracardiac injection in a mouse MI model is presented. This protocol will make modRNA delivery more accessible for basic and translational research.

This protocol presents a simple and coherent way to transiently upregulate a gene of interest using modRNA after myocardial infarction in mice.

Myocardial infarction (MI) is a leading cause of morbidity and mortality in the Western world. In the past decade, gene therapy has become a promising ...

Genetics

Isolation and Genetic Manipulation of Adult Cardiac Myocytes for Confocal Imaging

Cardiac myocytes isolated from adult hearts are widely accepted as a model somewhere half way between embryonic and neonatal muscle cells on one side and a working heart on the other. Thus, cardiomyocytes serve as good models for cardiac cellular physiology and pathophysiology, for pharmaceutical investigations as well as for the exploration of transgenic animal models. Here we describe a method of isolating the cells from the heart. Furthermore we show how a genetic manipulation on cardiac myocytes can be performed without breeding a transgenic animal: This is the combination of long term culture (1 week) and adenoviral gene transfer. The latter one is described from the construction of the virus to the transduction of the cells. It can be used for the expression of genetically encoded biosensors (GEBs), fluorescent fusion proteins, but also for protein over expression and down regulation, e.g. using RNAi. Here we provide an example for the expression of a fusion protein staining a subcellular structure (Golgi). Such protein expression can be visualized by confocal imaging of z-stacks for a 3D-reconstruction of subcellular structures. The protocol comprises state-of-the-art in cell culture, molecular biology and biophysics and thus provides an approach for exploring new horizons in cellular cardiology.

Adult cardiac myocytes are primary cells that can be isolated from animal hearts and cultured for several days. Within this culture period adenoviral gene transfer can be used to express genetically encoded biosensors (GEBs) or fluorescent fusion proteins. Both approaches allow cellular investigations by means of confocal microscopy.

Cardiac myocytes isolated from adult hearts are widely accepted as a model somewhere half way between embryonic and neonatal muscle cells on one side and ...

Ultrasound-guided Transthoracic Intramyocardial Injection in Mice

Murine models of cardiovascular disease are important for investigating pathophysiological mechanisms and exploring potential regenerative therapies. Experiments involving myocardial injection are currently performed by direct surgical access through a thoracotomy. While convenient when performed at the time of another experimental manipulation such as coronary artery ligation, the need for an invasive procedure for intramyocardial delivery limits potential experimental designs. With ever improving ultrasound resolution and advanced noninvasive imaging modalities, it is now feasible to routinely perform ultrasound-guided, percutaneous intramyocardial injection. This modality efficiently and reliably delivers agents to a targeted region of myocardium. Advantages of this technique include the avoidance of surgical morbidity, the facility to target regions of myocardium selectively under ultrasound guidance, and the opportunity to deliver injectate to the myocardium at multiple, predetermined time intervals. With practiced technique, complications from intramyocardial injection are rare, and mice quickly return to normal activity on recovery from anesthetic. Following the steps outlined in this protocol, the operator with basic echocardiography experience can quickly become competent in this versatile, minimally invasive technique.

Echocardiography-guided percutaneous intramyocardial injection represents an efficient, reliable, and targetable modality for the delivery of gene transfer agents or cells into the murine heart. Following the steps outlined in this protocol, the operator can quickly become competent in this versatile, minimally invasive technique.

Murine models of cardiovascular disease are important for investigating pathophysiological mechanisms and exploring potential regenerative therapies. ...

Medicine

Assessing Cardiac Reprogramming using High Content Imaging Analysis

The goal of this protocol is to describe a method for quantifying induced cardiomyocyte-like cells (iCMs), which are directly reprogrammed in vitro by a reprogramming technique. Cardiac reprogramming provides a strategy to generate new cardiomyocytes. By introducing core cardiogenic transcription factors into fibroblasts; fibroblasts can be converted to iCMs without transition through the pluripotent stem cell state. However, the conversion rate of fibroblasts to iCMs still remains low. Accordingly, there have been numerous additional approaches to enhance cardiac reprogramming efficiency. Most of these studies assessed cardiac reprogramming efficiency using flow cytometry, while at the same time performed immunocytochemistry to visualize iCMs. Thus, at least two separate sets of reprogramming experiments are required to demonstrate the success of iCM reprogramming. In contrast, automated high content imaging analysis will provide both quantification and qualification of iCM reprogramming with a relatively small number of cells. With this method, it is possible to directly assess the quantity and quality of iCMs with a single reprogramming experiment. This approach will be able to facilitate future cardiac reprogramming studies that require large-scale reprogramming experiments such as screening genetic or pharmacological factors for enhancing reprogramming efficiency. In addition, the application of high content imaging analysis protocol is not limited to cardiac reprogramming. It can be applied to reprogramming of other cell lineages as well as any immunostaining experiments which need both quantification and visualization of immunostained cells.

We present a protocol to quantify directly reprogrammed induced cardiomyocyte-like cells (iCMs) in vitro using high content imaging analysis. This method allows us to quantify the efficiency of cardiac reprogramming in an automated manner and to directly visualize iCMs.

The goal of this protocol is to describe a method for quantifying induced cardiomyocyte-like cells (iCMs), which are directly reprogrammed in vitro by a ...

Developmental Biology

In Vivo Two-Color 2-Photon Imaging of Genetically-Tagged Reporter Cells in the Skin

Fibroblasts are mesenchymal cells that change their morphology upon activation, ultimately influencing the microenvironment of the tissue they are located in. Although traditional imaging techniques are useful in identifying protein interactions and morphology in fixed tissue, they are not able to give insight as to how quickly cells are able to bind and uptake proteins, and once activated how their morphology changes in vivo. In the present study, we ask 2 major questions: 1) what is the time-course of fibroblast activation via toll-like receptor-4 (TLR4) and lipopolysaccharide (LPS) interaction and 2) how do these cells behave once activated? Using 2-photon imaging, we have developed a novel technique to assess the ability of LPS-FITC to bind to its cognate receptor, TLR4, expressed on peripheral fibroblasts in the genetic reporter mouse line; FSP1cre; tdTomatolox-stop-lox&#160;in vivo. This unique approach allows researchers to create in-depth, time-lapse videos and/or pictures of proteins interacting with live cells that allows one to have an increased level of granularity in understanding how proteins can alter cellular behavior.

Morphological changes occur in immune responsive fibroblast cells following activation and promote alterations in cellular recruitment. Utilizing 2-photon imaging in conjunction with a genetically engineered Fibroblast-specific protein 1 (FSP1)-cre; tdTomato floxed-stop-floxed (TB/TB) mouse line and green fluorescently tagged lipopolysaccharide-FITC, we can illustrate highly specific uptake of lipopolysaccharide in dermal fibroblasts and morphological changes in vivo.

Fibroblasts are mesenchymal cells that change their morphology upon activation, ultimately influencing the microenvironment of the tissue they are located ...

Bioengineering

Percutaneous Contrast Echocardiography-guided Intramyocardial Injection and Cell Delivery in a Large Preclinical Model

Cell and gene therapy are exciting and promising strategies for the purpose of cardiac regeneration in the setting of heart failure with reduced ejection fraction (HFrEF). Before they can be considered for use, and implemented in humans, extensive preclinical studies are required in large animal models to evaluate the safety, efficacy, and fate of the injectate (e.g., stem cells) once delivered into the myocardium. Small rodent models offer advantages (e.g., cost effectiveness, amenability for genetic manipulation); however, given inherent limitations of these models, the findings in these rarely translate into the clinic. Conversely, large animal models such as rabbits, have advantages (e.g., similar cardiac electrophysiology compared to humans and other large animals), whilst retaining a good cost-effective balance. Here, we demonstrate how to perform a percutaneous contrast echocardiography-guided intramyocardial injection (IMI) technique, which is minimally invasive, safe, well tolerated, and very effective in the targeted delivery of injectates, including cells, into several locations within the myocardium of a rabbit model. For the implementation of this technique, we also have taken advantage of a widely available clinical echocardiography system. After putting in practice the protocol described here, a researcher with basic ultrasound knowledge will become competent in the performance of this versatile and minimally invasive technique for routine use in experiments, aimed at hypothesis testing of the capabilities of cardiac regenerative therapeutics in the rabbit model. Once competency is achieved, the whole procedure can be performed within 25 min after anaesthetizing the rabbit.

Novel therapeutic strategies in cardiac regenerative medicine require extensive and detailed studies in large preclinical animal models before they can be considered for use in humans. Here, we demonstrate a percutaneous contrast echocardiography-guided intramyocardial injection technique in rabbits, which is valuable for hypothesis testing the efficacy of such novel therapies.

Cell and gene therapy are exciting and promising strategies for the purpose of cardiac regeneration in the setting of heart failure with reduced ejection ...

Live Cell Imaging of Primary Rat Neonatal Cardiomyocytes Following Adenoviral and Lentiviral Transduction Using Confocal Spinning Disk Microscopy

Primary rat neonatal cardiomyocytes are useful in basic in vitro cardiovascular research because they can be easily isolated in large numbers in a single procedure. Due to advances in microscope technology it is relatively easy to capture live cell images for the purpose of investigating cellular events in real time with minimal concern regarding phototoxicity to the cells. This protocol describes how to take live cell timelapse images of primary rat neonatal cardiomyocytes using a confocal spinning disk microscope following lentiviral and adenoviral transduction to modulate properties of the cell. The application of two different types of viruses makes it easier to achieve an appropriate transduction rate and expression levels for two different genes. Well focused live cell images can be obtained using the microscope&#8217;s autofocus system, which maintains stable focus for long time periods. Applying this method, the functions of exogenously engineered proteins expressed in cultured primary cells can be analyzed. Additionally, this system can be used to examine the functions of genes through the use of siRNAs as well as of chemical modulators.

This protocol describes a method of live cell imaging using primary rat neonatal cardiomyocytes following lentiviral and adenoviral transduction using confocal spinning disk microscopy. This enables detailed observations of cellular processes in living cardiomyocytes.

Primary rat neonatal cardiomyocytes are useful in basic in vitro cardiovascular research because they can be easily isolated in large numbers in a single ...

Large Animal Model for Evaluating the Efficacy of the Gene Therapy in Ischemic Heart

Coronary artery disease is one of the significant causes of mortality and morbidity worldwide. Despite the progression of current therapeutics, a considerable proportion of coronary artery disease patients remain symptomatic. Gene therapy-mediated therapeutic angiogenesis offers a novel therapeutic method for improving myocardial perfusion and relieving symptoms. Gene therapy with different angiogenic factors has been studied in few clinical trials. Due to the novelty of the method, the progress of myocardial gene therapy is a continuous path from bench to bedside. Therefore, large animal models are needed for evaluating the safety and efficacy. The more the large animal model identifies the original disease and the endpoints used in clinics, the more predictable outcomes are from clinical trials. Here, we introduce a large animal model for evaluating the efficacy of the gene therapy in the ischemic porcine heart. We use clinically relevant imaging methods such as ultrasound imaging and 15H2O-PET. For targeting the gene transfers into the desired area, electroanatomical mapping is used. The aim of this method is: (1) to mimic chronic coronary artery disease, (2) to induce therapeutic angiogenesis at hypoxic areas of the heart, and (3) to evaluate the safety and efficacy of the gene therapy by using relevant endpoints.

Myocardial gene therapy for ischemic heart disease holds great promise for future therapeutics. Here, we introduce a large animal model for evaluating the efficacy of gene therapy in the ischemic heart.

Coronary artery disease is one of the significant causes of mortality and morbidity worldwide. Despite the progression of current therapeutics, a ...

Live Cell Imaging and 3D Analysis of Angiotensin Receptor Type 1a Trafficking in Transfected Human Embryonic Kidney Cells Using Confocal Microscopy

Live-cell imaging is used to simultaneously capture time-lapse images of angiotensin type 1a receptors (AT1aR) and intracellular compartments in transfected human embryonic kidney-293 (HEK) cells following stimulation with angiotensin II (Ang II). HEK cells are transiently transfected with plasmid DNA containing AT1aR tagged with enhanced green fluorescent protein (EGFP). Lysosomes are identified with a red fluorescent dye. Live-cell images are captured on a laser scanning confocal microscope after Ang II stimulation and analyzed by software in three dimensions (3D, voxels) over time. Live-cell imaging enables investigations into receptor trafficking and avoids confounds associated with fixation, and in particular, the loss or artefactual displacement of EGFP-tagged membrane receptors. Thus, as individual cells are tracked through time, the subcellular localization of receptors can be imaged and measured. Images must be acquired sufficiently rapidly to capture rapid vesicle movement. Yet, at faster imaging speeds, the number of photons collected is reduced. Compromises must also be made in the selection of imaging parameters like voxel size in order to gain imaging speed. Significant applications of live-cell imaging are to study protein trafficking, migration, proliferation, cell cycle, apoptosis, autophagy and protein-protein interaction and dynamics, to name but a few.

Here we present a protocol to image cells expressing green fluorescent protein-tagged angiotensin type 1a receptors during endocytosis initiated by angiotensin II treatment. This technique includes labeling lysosomes with a second fluorescent marker, and then utilizing software to analyze the co-localization of receptor and lysosome in three dimensions over time.

Live-cell imaging is used to simultaneously capture time-lapse images of angiotensin type 1a receptors (AT1aR) and intracellular compartments in ...

Delivery of Cardioactive Therapeutics in a Porcine Myocardial Infarction Model

Myocardial infarction is one of the leading causes of death and disability worldwide, and there is an urgent need for novel cardioprotective or regenerative strategies. An essential component of drug development is determining how a novel therapeutic is to be administered. Physiologically relevant large animal models are of critical importance in assessing the feasibility and efficacy of various therapeutic delivery strategies. Due to their similarities to humans in cardiovascular physiology, coronary vascular anatomy, and heart weight to body weight ratio, swine is one of the preferred species in the preclinical evaluation of new therapies for myocardial infarction. The present protocol describes three methods of administering cardioactive therapeutic agents in a porcine model. After percutaneously induced myocardial infarction, female landrace swine received treatment with novel agents through either: (1) thoracotomy and transepicardial injection, (2) catheter-based transendocardial injection, or (3) intravenous infusion via jugular vein osmotic minipump. The procedures employed for each technique are reproducible, resulting in reliable cardioactive drug delivery. These models can be easily adapted to suit individual study designs, and each of these delivery techniques can be used to investigate a variety of possible interventions. Therefore, these methods are a useful tool for translational scientists pursuing novel biological approaches in cardiac repair following myocardial infarction.

The present protocol describes three methods of administering cardioactive therapeutic agents in a porcine model. Female landrace swine received treatment through either: (1) thoracotomy and transepicardial injection, (2) catheter-based transendocardial injection, or (3) intravenous infusion via jugular vein osmotic minipump.