The authors would like to acknowledge Michael Solga and the rest of the Flow Cytometry Core staff for providing flow cytometry training and services. The authors would also like to acknowledge Shiva Sai Krishna Dasa, Dustin K. Bauknight, Melissa A. Marshall, James C. Garmey, Chantel McSkimming, Aditi Upadhye, and Prasad Srikakulapu for their assistance with liposome preparation (SSKD, DKB), tissue harvests (MAM, JCG), and flow cytometry staining and sample acquisition (AU, PS, CM). This work was supported by AstraZeneca, R01HL 136098, R01HL 141123 and R01HL 148109, AHA 16PRE30770007, and T32 HL007284 grants.

<ol><li>Allen, T. M., Cullis, P. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Liposomal+drug+delivery+systems%3A+from+concept+to+clinical+applications%5BTitle%5D)+AND+%22Advanced+Drug+Delivery+Reviews%22%5BJournal%5D)">Liposomal drug delivery systems: from concept to clinical applications.</a> Advanced Drug Delivery Reviews. 65 (1), 36-48 (2013).</li>
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<li>Dasa, S. S. K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Plectin-targeted+liposomes+enhance+the+therapeutic+efficacy+of+a+PARP+inhibitor+in+the+treatment+of+ovarian+cancer%5BTitle%5D)+AND+%22Theranostics%22%5BJournal%5D)">Plectin-targeted liposomes enhance the therapeutic efficacy of a PARP inhibitor in the treatment of ovarian cancer.</a> Theranostics. , (2018).</li>
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The authors have nothing to disclose.

Here we describe a three-part protocol to (i) prepare liposomes that are labeled with a fluorescent lipid dye and loaded with an anti-diabetic compound, tesaglitazar, (ii) administer liposomes to a mouse via retro-orbital injection, and (iii) harvest, process, and stain tissues to detect liposome uptake at a cellular level by flow cytometry. This protocol reviews preparation of approximately 150-&#181;m liposomes and assessment of uptake in adipose, blood, and the spleen. The liposome preparation is scalable, performed mostly at room temperature, and utilizes reverse-phase evaporation to maximize drug loading and removal of organic solvents. Using this protocol, up to 2 mg/mL tesaglitazar concentration can be achieved in the purified liposome sample. The prepared liposomes can be stored in HEPES buffer at 4 &#176;C for over a year. In our experience, they demonstrated minimal variation of mean particle size. Under 10% of drug content loss was demonstrated spectrophotometrically, following ultrafiltration separation of liposomes from external drug with a 10 kDa centrifugal filter.
During liposome preparation, there are some critical steps and factors to consider. First, the order of the protocol steps is important and must be adhered to. Second, the pH of the solution used when loading tesaglitazar must be maintained at 7.4 in order to maximize solubility and effective loading. Third, proper assembly of equipment and filters ensures that the output of each step is of the proper size and purity. For example, if 100- and 200-nm filters are not assembled properly, a more heterogenous and improperly-sized batch of liposomes may result. Fourth, complete removal of Ca-acetate prior to drug-loading is needed to maximize the transfer of tesaglitazar into the liposomes. To test for complete removal of Ca-acetate, use high-speed sedimentation to remove the liposomes and then measure Ca-acetate levels in the non-liposomal solution. Fifth, it is important to weigh and record the mass of all materials added to the liposome preparation at each step. This ensures that proper concentrations can be calculated and needed ratios of materials are maintained. Finally, if the technique is not properly executed, there may be an undesirable level of heterogeneity. It is important to thoroughly check this parameter using DLS and other approaches such as electron microscopy. To improve homogeneity, consider adjusting the selected filter size or stacking two filters.
Additionally, it is critical that controls and an antibody panel for flow cytometry are planned and optimized prior to conducting this protocol in full (Table 1, Table 3). Antibodies should be tested to ensure proper concentrations are used for staining and that overlap between fluorophores is minimal. The excitation and emission of the dye used during liposome preparation must also be factored into panel planning. In our results, we utilized DiD, which has a similar excitation and emission to fluorophores such as Allophycocyanin (APC) and AlexaFluor 647. Thus, we did not select antibodies conjugated to these fluorophores in our antibody panel. Furthermore, isotype controls are not included in this protocol. This is because the antibodies selected for this protocol are well-validated, commercially available antibodies. However, if interested in using an antibody that has not been optimized previously, please consider testing the antibody against an isotype control on the tissues of interest prior to conducting the full experiment.
While this protocol demonstrates how to extract and process the blood, spleen, inguinal adipose, and epididymal adipose tissues from the mouse post-treatment, this general approach can be applied to other tissues. Depending on the tissue of interest, processing and digestion protocols may need to be altered as is published for the following tissues: lung21, liver22, peritoneal cavity3, bone marrow3,23, brain24.
An important limitation of this method to consider is that uptake can only be assessed at one time point per animal. Thus, it may be advantageous to couple this protocol with other non-invasive imaging techniques or plan accordingly to ensure sufficient resources for conducting the assessment. Timing of cellular uptake and cellular turn over are important factors to consider: liposomes will circulate throughout the body in the first 24 h and depending on the lifespan of the cells that take up liposomes or how they respond to uptake, cell death or further phagocytosis may occur. Our previous study demonstrated changes in the population characteristics of DiD+ populations at different time points3. For that reason, evaluating uptake at earlier time points or time points most relevant to the biology of mechanism of interest is important. Additionally, while quantification of cell uptake in the entire tissue can be performed with this protocol, flow cytometry cannot reveal tissue localization. Coupling this approach with histological methods can help to address this limitation.
In general, this protocol complements existing methodology such as histology and whole-body fluorescence imaging. With the continued advancements in flow cytometry tools and methods, the development of larger panels to more and more specific cell populations will become possible. We suggest that this protocol be used in addition to the aforementioned methods as this will improve the evaluation of cellular uptake and also provide the opportunity to validate the outcomes observed by flow cytometry. For example, should it be found that a majority of the particles in adipose tissue were taken up by macrophages by flow cytometry. Immunofluorescence of an additional aliquot of the same adipose tissue could be saved, fixed, sectioned, and stained for macrophage markers to verify that the cell type does indeed take up liposomes. This approach should add rigor to nanoparticle biodistribution assays conducted: validating cell-specific targeting, quantifying cellular uptake, identifying off-target uptake, and hopefully providing information to generate mechanistic hypotheses for observed therapeutic outcomes. This protocol can be also be adapted for future studies using different liposomes, investigating uptake in other tissues, and testing new compounds in the setting of obesity and dysmetabolism or any other disease in which nanoparticle-delivery is a feasible therapeutic option.

As interest in developing therapies utilizing nanoparticles drug delivery grows, the methods to prepare and assess particle distribution and uptake must continue to advance, expand, and be accessible to the research community1,2. This protocol was developed to assess the exact cell types that took up liposomes in vivo following a treatment with DiD-labeled liposomes loaded with tesaglitazar, a peroxisome proliferator-activated receptor (PPAR)-&#945;/&#947; agonist3,4. In these studies, we were able to assess which cell types were directly impacted by liposomal tesaglitazar treatment, the efficacy of targeting moieties, and generate hypotheses to explain the treatment outcomes we observed. Furthermore, established biological functions in a variety of cell types suggest that phagocytic cells such as macrophages, dendritic cells, and liver-specific Kupffer cells take up most of the liposomes5,6,7. Utilizing this protocol, we have demonstrated that non-classical phagocytes could also take up liposomes3,4.
This protocol presents an optimized method for solubilizing tesaglitazar, preparing liposomes by reverse-phase evaporation, and using calcium acetate as an attractant for remote drug loading. The methods presented are accessible to many labs and lack hard-to-acquire materials and steps requiring high temperatures. The protocol produces liposomes of sizes which are optimal for increased circulation in vivo8. Furthermore, as summarized by Su et al., to date, methods to evaluate in vivo liposome distribution and tissue uptake have been studied and tested in depth9. Positron emission tomography (PET), magnetic resonance imaging (MRI) and fluorescence molecular tomography (FMT) methods are applied to quantify tissue-specific biodistribution and uptake9,10,11. While these methods have been optimized to maximize detection in vivo, they still lack the ability to quantify liposome uptake in vivo at cellular resolution. The protocol presented here aims to accomplish this need through the use of flow cytometry. Finally, for this protocol, cellular uptake was narrowed down to a few tissues including adipose tissue. There is a growing body of literature investigating the potential for use of nanoparticles to deliver therapies in the setting of obesity, dysmetabolism, and inflammation12,13,14,15,16,17. As such, we felt it important to share a protocol with effective methods for processing and analyzing adipose tissue&#8212;one of the tissues that plays an important role in these pathologies.

<table><tbody><tr><td>1-mL syringe</td><td>BD</td><td>309659</td><td></td></tr><tr><td>10-mL syringe</td><td>BD</td><td>302995</td><td></td></tr><tr><td>25-gauge needle, sterile for retro-orbital injection</td><td>BD</td><td>305122</td><td></td></tr><tr><td>27-gauge needle, sterile for retro-orbital injection</td><td>BD</td><td>305620</td><td></td></tr><tr><td>Anti-mouse B220 BV421</td><td>Biolegend</td><td>103251</td><td>Clone RA3-6B2</td></tr><tr><td>Anti-mouse CD115 PE</td><td>eBioscience</td><td>12-1152-82</td><td>Clone AFS98</td></tr><tr><td>Anti-mouse CD11b PerCP Cy5.5 antibody</td><td>BD Biosciences</td><td>550993</td><td>Clone M1/70</td></tr><tr><td>Anti-mouse CD11c APC ef780 antibody</td><td>eBioscience</td><td>47-0114-82</td><td>Clone N418</td></tr><tr><td>Anti-mouse CD19 PE CF594</td><td>BD Biosciences</td><td>562291</td><td>Clone 1D3</td></tr><tr><td>Anti-mouse CD3 FITC antibody</td><td>BD Biosciences</td><td>553061</td><td>Clone 145-2C11</td></tr><tr><td>Anti-mouse CD31 BV605</td><td>Biolegend</td><td>102427</td><td>Clone 390</td></tr><tr><td>Anti-mouse CD45 PerCP</td><td>BD Biosciences</td><td>557235</td><td>Clone 30-F11</td></tr><tr><td>Anti-mouse F4/80 PE Cy7</td><td>Biolegend</td><td>123114</td><td>Clone BM8</td></tr><tr><td>Bovine serum albumin</td><td>Gemini Bio-products</td><td>700-107P</td><td></td></tr><tr><td>Desalting spin-column</td><td>ThermoFisher</td><td>89889, 89890</td><td>Zeba spin column</td></tr><tr><td>DPBS</td><td>Gibco</td><td>14190-144</td><td></td></tr><tr><td>Dynamic Light Scattering, Nicomp 370</td><td>Particle Sizing System, Inc</td><td></td><td></td></tr><tr><td>FIX &amp;amp; PERM Cell Permeabilization Kit</td><td>ThermoFisher Scientific</td><td>GAS004</td><td></td></tr><tr><td>Gauze sponges</td><td>Dermacea</td><td>441211</td><td></td></tr><tr><td>Heparin</td><td>Sigma</td><td>3393-1MU</td><td></td></tr><tr><td>Liposome extruder</td><td>Millipore Sigma</td><td>Z373400</td><td>LiposoFast</td></tr><tr><td>Live/Dead Aqua</td><td>ThermoFisher Scientific</td><td>L34957</td><td></td></tr><tr><td>Nanosight</td><td>Malvern Instruments Ltd</td><td>NS300</td><td></td></tr><tr><td>Ophthalmic lubricant</td><td>Optixcare</td><td></td><td>20g/70 oz Sterile</td></tr><tr><td>Paraformaldehyde, 16% w/v aq. soln., methanol free</td><td>Alfa Aesar</td><td>433689L</td><td></td></tr><tr><td>Polyethylene vial for mincing</td><td>Wheaton</td><td>986701</td><td></td></tr><tr><td>Rotary evaporator</td><td>Buchi</td><td>Re111</td><td></td></tr><tr><td>Sonicator</td><td>Misonix</td><td>XL2020</td><td></td></tr><tr><td>T/Pump Heat therapy pump and pad</td><td>Gaymer Industries</td><td>TP-500</td><td></td></tr><tr><td>Tesaglitazar</td><td>Tocris</td><td>3965</td><td></td></tr><tr><td>Track-etched polycarbonate membranes</td><td>Thomas Scientific</td><td>1141Z**</td><td>Whatman, Nuclepore Polycarbonate hydrophilic membranes</td></tr><tr><td>ZetaSizer/DLS-ELS system</td><td>Malvern Instruments Ltd</td><td></td><td></td></tr></tbody></table>

preparation, administration, and assessment of in vivo tissue-specific cellular uptake of fluorescent dye-labeled liposomes

There is a growing interest in using liposomes to deliver compounds in vivo particularly for targeted treatment approaches. Depending on the liposome formulation, liposomes may be preferentially taken up by different cell types in the body. This may influence the efficacy of the therapeutic particle as progression of different diseases is tissue- and cell-type-specific. In this protocol, we present one method for synthesizing and fluorescently labeling liposomes using DSPC, cholesterol, and PEG-2000 DSPE and the lipid dye DiD as a fluorescent label. This protocol also presents an approach for delivering liposomes in vivo and assessing cell-specific uptake of liposomes using flow cytometry. This approach can be used to determine the types of cells that take up liposomes and quantify the distribution and proportion of liposome-uptake across cell types and tissues. While not mentioned in this protocol, additional assays such as immunofluorescence and single-cell fluorescence imaging on a cytometer will strengthen any findings or conclusions made as they permit assessment of intracellular staining. Protocols may also need to be adapted depending on the tissue(s) of interest.

Liposome Production
Results published here are similar to those in our previously published work3,4,20. Utilizing the protocol presented here, we expect to produce liposomes of approximately 150&#8210;160 nm in size. DLS reveals an average liposome diameter of 163.2 nm and a zeta potential of -19.2 mV (Figure 1A). Cryogenic electron microscopy (cryo-EM) imaging reveals circular liposomes (Figure 1B) and the DLS diagram reveals a relatively small standard deviation from the average diameter (Figure 1C).
Positive liposome binding requires a PBS-treated control
Prior studies from our group employing this protocol investigated what cell subsets in adipose SVF, spleen, and blood bound to liposomes following one week of in vivo administration3,4. Using a PBS-treated mouse, peritoneal cavity and spleen cells were stained with the same antibody panel used on samples from liposome-treated mice. Tissues were harvested after one week of treatments (Figure 2A). The samples from the PBS-treated mouse served as a DiD FMO with which to create positive DiD gates (Figure 2B,C). A positive gate can be created using DiD-positive signal, but samples lacking DiD signal must also be used to verify that the positive gate does not include any DiD-negative samples.
Titrations are needed to optimize fluorescence signals
Prior to executing a full experiment, various conditions including the concentration of fluorescently conjugated antibodies used during cell staining and of lipid dye used during liposome preparation must be optimized. Flow cytometers have an upper limit of detection for fluorescence intensity, so too much dye incorporated in the liposomes will lead to unquantifiable levels of DiD signal in samples run through the cytometer. Furthermore, too much DiD in the liposomes may lead to high levels of non-specific dye transfer, which could skew cellular uptake results. Figure 3 reports results from an experiment in which concentrations of lipid dye were titrated to identify the concentration that would produce an optimal signal within the detection range of the flow cytometer that was used. This was conducted on the tissues of interest for the final experiment: Blood (Figure 3A), inguinal adipose SVF (Figure 3B), and epididymal adipose SVF (Figure 3C). The concentrations selected for testing were 10 mg of DiD (High, red), 1 mg DiD (Middle, blue), or 0.1 mg of DiD (Low, grey) per 1 mL of liposomes. The highest concentration used in the liposomes was too high and surpassed the quantifiable range of the cytometer in all three tissues (Figure 3A&#8210;C, red). The lowest concentration of DiD showed some signal (Figure 3A&#8210;C, grey), but a clear population beyond the PBS-treated cells (Figure 3A&#8210;C, black) was not observed. When quantified, the arithmetic mean of the DiD MFI for each tissue and concentration demonstrated a clear distinction between PBS controls and the middle concentration of DiD (Figure 3D). Thus, as indicated in the protocol, we selected the middle concentration (Figure 3, blue) to use in our liposome preparation.
The use of multi-antibody panel allows for identification of liposome uptake by different cell subsets
Using the panel outlined in Table 3, cells were stained with antibodies against markers for a macrophages, B cells, T cells, dendritic cells, monocytes, and endothelial cells (Figure 4). Slightly different gating strategies are required for each tissue type, but most of the same cell types can be identified in each. Some exceptions include endothelial cells, which are not normally found in the blood, and monocytes, which are typically at higher frequency in the blood than other tissues. Once populations are identified, total size of each cell population and the frequency at which they are DiD+ can be quantified. Further calculations can be performed to characterize the DiD+ population: what percent of DiD+ cells are macrophages, endothelial cells, etc. Please note, these are example gating strategies, but not the only way to analyze the samples. Analysis will be dictated by the selected panel and flow cytometer(s) available.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/61585/61585fig01.jpg" /> 
Figure 1: Example characteristics of prepared liposomes. 
(A) The size and zeta potential were measured as described above and have been reported in table form. Each parameter is presented as the mean &#177; the standard deviation. (B) Cryo-EM was used to image the prepared liposomes. The white scale bar is 50 nm in length. (C) DLS was used to generate a histogram of the diameter of liposomes in this prep. This figure is adapted from Osinski et al.3. <a href="https://www.jove.com/files/ftp_upload/61585/61585fig01large.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/61585/61585fig02.jpg" /> 
Figure 2: Representative DiD staining from PBS- or Liposome-treated mice. 
(A) Experimental schematic for PBS and liposome treatments. PBS or liposomes were injected three times over the course of one week. Tissues were harvested on Day 8 of treatment. (B, C) Representative flow plots reveal positive DiD staining in liposome-treated (C), but not PBS-treated (B) mice. FSC, forward scatter. <a href="https://www.jove.com/files/ftp_upload/61585/61585fig02large.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/61585/61585fig03.jpg" /> 
Figure 3: Titration of DiD in liposomes. 
Liposomes were prepared with three different concentrations of DiD and injected into mice. Grey indicates the low concentration at 0.1 mg DiD per 1 mL of liposomes, blue indicates the middle concentration at 1 mg DiD/mL liposomes, and red indicates the high concentration at 10 mg DiD/mL liposomes. A PBS-treated mouse was used as a negative control (black). Blood (A, circle), inguinal adipose (B, triange), and epididymal adipose (C, square) were harvested 24 h post-injection and processed to isolate a single-cell suspension. These samples were run on a flow cytometer to the level of detectable DiD. Tissue-specific histograms with overlays of each treatment group are presented to demonstrate fluorescence intensity per concentration (A&#8210;C). The arithmetic mean of DiD was also quantified for each tissue and concentration and plotted (D). SSC = side scatter. <a href="https://www.jove.com/files/ftp_upload/61585/61585fig03large.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/61585/61585fig04v4.jpg" /> 
Figure 4: Representative flow cytometry analysis of cell subsets in adipose SVF, blood, and spleen. 
(A&#8210;C) Schematic representative of gating strategy to identify cell subsets and DiD+ cells in adipose SVF (A), spleen (B), and blood (C). Abbreviations: FSC = forward scatter; LD = live/dead; L-DCs = lymphoid dendritic cells; M-DCs = myeloid dendritic cells; SSC = side scatter. This figure is adapted from Osinski et al.3. <a href="https://www.jove.com/files/ftp_upload/61585/61585fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Control</td>
			<td>Purpose</td>
		</tr>
		<tr>
			<td rowspan="5">Mouse treated with PBS or saline</td>
			<td>Use the cells from this mouse for the following flow cytometry controls:</td>
		</tr>
		<tr>
			<td>1. Unstained cells</td>
		</tr>
		<tr>
			<td>2. Live/dead single stain</td>
		</tr>
		<tr>
			<td>3. Cells stained with the full panel, but lacking the liposome fluorescence to determine positive liposome signal during analysis</td>
		</tr>
		<tr>
			<td>This/these mouse/mice will also be used to determine if liposomes have any effects in vivo as you will have a non-liposome control in your experiment.</td>
		</tr>
		<tr>
			<td>Unloaded liposomes</td>
			<td>If you are loading a compound in your liposomes, a portion of your liposome batch should be synthesized without the compound. This accounts for any in vivo effects of the liposomes alone.</td>
		</tr>
		<tr>
			<td>DiD alone</td>
			<td>Since DiD can also be taken up by cellular membranes, allocating some mice to receive free dye at an amount equal to that found in the liposomes will help account for any background membrane staining.</td>
		</tr>
		<tr>
			<td>Fluorescence-minus-one (FMO) controls</td>
			<td>These are cells stained with all but one of the antibodies in your panel. Like #3 in the box above, this aids in determining true positive signal for that antibody during analysis</td>
		</tr>
	</tbody>
</table>
Table 1: Controls to use in this protocol.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Solution</td>
			<td>Components</td>
			<td>Approximate volume needed per batch/mouse</td>
		</tr>
		<tr>
			<td colspan="3">Liposome preparation</td>
		</tr>
		<tr>
			<td>Calcium acetate</td>
			<td>1 M calcium acetate in H2O</td>
			<td>50 mL</td>
		</tr>
		<tr>
			<td>HEPES buffer</td>
			<td>10 mM HEPES in H2O, pH 7.4</td>
			<td>50 mL</td>
		</tr>
		<tr>
			<td>Tesaglitazar in HEPES</td>
			<td>in 10 mM HEPES</td>
			<td>10 mL</td>
		</tr>
		<tr>
			<td colspan="3">Tissue harvest, processing, and staining</td>
		</tr>
		<tr>
			<td>Phosphate-buffered solution (PBS)</td>
			<td>137 mM NaCl, 2.7 mM KCl, 10 mM Na2HPO4, 1.8 mM KH2PO4 in distilled H2O</td>
			<td>2 mL</td>
		</tr>
		<tr>
			<td>PBS-Heparin</td>
			<td>0.1 mM Heparin in PBS</td>
			<td>10 mL</td>
		</tr>
		<tr>
			<td>HEPES buffer</td>
			<td>20 mM HEPES in PBS</td>
			<td>5 mL</td>
		</tr>
		<tr>
			<td>Digestion buffer</td>
			<td>2 mg/mL Collagenase Type I in HEPES buffer</td>
			<td>5 mL</td>
		</tr>
		<tr>
			<td>AKC lysis buffer</td>
			<td>0.158 M NH3Cl, 10 mM KHCO3, 0.1 mM Na2EDTA in ddH2O, pH 7.2</td>
			<td>15 mL</td>
		</tr>
		<tr>
			<td>FACS buffer</td>
			<td>1% BSA, 0.05% NaN3 in PBS</td>
			<td>15 mL</td>
		</tr>
		<tr>
			<td>Fc Block (diluted)</td>
			<td>1:50 Fc Block in FACS buffer</td>
			<td>250 &#181;L</td>
		</tr>
		<tr>
			<td>Fixation Buffer</td>
			<td>2% paraformaldehyde in PBS</td>
			<td>200 &#181;L</td>
		</tr>
	</tbody>
</table>
Table 2: Solutions to prepare.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>A</td>
			<td>B</td>
			<td>C</td>
			<td>D</td>
		</tr>
		<tr>
			<td colspan="4">Extracellular Staining (2x antibody mix)</td>
		</tr>
		<tr>
			<td>Antigen</td>
			<td>Fluorophore</td>
			<td>Ab volume per 100 &#181;L test</td>
			<td>Total volume needed:</td>
		</tr>
		<tr>
			<td>CD45</td>
			<td>PerCP</td>
			<td>0.5 &#181;L</td>
			<td>Column C x 1.2 x Total # samples</td>
		</tr>
		<tr>
			<td>CD11b</td>
			<td>PerCP Cy5.5</td>
			<td>0.25 &#181;L</td>
			<td>(0.5 &#181;L/test) x (1.2) x (# samples)</td>
		</tr>
		<tr>
			<td>F4/80</td>
			<td>PE Cy7</td>
			<td>0.25 &#181;L</td>
			<td>(0.25 &#181;L/test) x (1.2) x (# samples)</td>
		</tr>
		<tr>
			<td>CD19</td>
			<td>PE-CF594</td>
			<td>1 &#181;L</td>
			<td>(0.25 &#181;L/test) x (1.2) x (# samples)</td>
		</tr>
		<tr>
			<td>CD3</td>
			<td>FITC</td>
			<td>1 &#181;L</td>
			<td>(1.0 &#181;L/test) x (1.2) x (# samples)</td>
		</tr>
		<tr>
			<td>CD31</td>
			<td>BV605</td>
			<td>0.25 &#181;L</td>
			<td>etc&#8230;</td>
		</tr>
		<tr>
			<td>CD11c</td>
			<td>APC ef780</td>
			<td>1 &#181;L</td>
			<td></td>
		</tr>
		<tr>
			<td>CD115</td>
			<td>PE</td>
			<td>1.5 &#181;L</td>
			<td></td>
		</tr>
		<tr>
			<td colspan="4">To create your antibody mix, combine the antibodies calculated in column D with FACS buffer or Brilliant Violet Staining Buffer* to a final volume of (50 &#181;L x 1.2 x Total # samples)</td>
		</tr>
		<tr>
			<td colspan="4">Live/Dead staining (1x)</td>
		</tr>
		<tr>
			<td>Live/Dead</td>
			<td>Fluorophore</td>
			<td>L/D volume per 200 uL test</td>
			<td>Total volume needed:</td>
		</tr>
		<tr>
			<td>Live/Dead</td>
			<td>Aqua</td>
			<td>0.67 &#181;L</td>
			<td>Column C x 1.2 x Total # samples</td>
		</tr>
		<tr>
			<td colspan="4">Intracellular Staining (1x)</td>
		</tr>
		<tr>
			<td>Antigen</td>
			<td>Fluorophore</td>
			<td>Ab volume per 50 &#181;L test</td>
			<td>Total volume needed:</td>
		</tr>
		<tr>
			<td>&#945;SMA</td>
			<td>FITC</td>
			<td>0.125</td>
			<td>Column C x 1.2 x Total # samples</td>
		</tr>
		<tr>
			<td colspan="4">*Brilliant Violet Staining Buffer should be used if more than one antibody conjugated to a Brilliant Violet fluorophore is being used in your panel.</td>
		</tr>
	</tbody>
</table>
Table 3: Example antibody panel and calculations of staining mixes to use for flow staining.

Watch this Scientific Journal Video about Preparation, Administration, and Assessment of In Vivo Tissue-Specific Cellular Uptake of Fluorescent Dye-Labeled Liposomes at JoVE.com

Preparation, Administration, and Assessment of In Vivo Tissue-Specific Cellular Uptake of Fluorescent Dye-Labeled Liposomes

The goal of this protocol is to synthesize fluorescently-labeled liposomes and use flow cytometry to identify in vivo localization of liposomes at a cellular level.

There is a growing interest in using liposomes to deliver compounds in vivo particularly for targeted treatment approaches. Depending on the liposome ...

preparation-administration-assessment-vivo-tissue-specific-cellular

Research

JoVE Journal

Bioengineering

4.2K Views.  University of Virginia. Het doel van dit protocol is om fluorescerend gelabelde liposomen te synthetiseren en flowcytometrie te gebruiken om in vivo lokalisatie van liposomen op cellulair niveau te identificeren.

Video: Preparation, Administration, and Assessment of In Vivo Tissue-Specific Cellular Uptake of Fluorescent Dye-Labeled Liposomes

A Comprehensive Procedure to Evaluate the In Vivo Performance of Cancer Nanomedicines

Inspired by the success of previous cancer nanomedicines in the clinic, researchers have generated a large number of novel formulations in the past decade. However, only a small number of nanomedicines have been approved for clinical use, whereas the majority of nanomedicines under clinical development have produced disappointing results. One major obstacle to the successful clinical translation of new cancer nanomedicines is the lack of an accurate understanding of their in vivo performance. This article features a rigorous procedure to characterize the in vivo behavior of nanomedicines in tumor-bearing mice at systemic, tissue, single-cell, and subcellular levels via the integration of positron emission tomography-computed tomography (PET-CT), radioactivity quantification methods, flow cytometry, and fluorescence microscopy. Using this approach, researchers can accurately evaluate novel nanoscale formulations in relevant mouse models of cancer. These protocols may have the ability to identify the most promising cancer nanomedicines with high translational potential or to aid in the optimization of cancer nanomedicines for future translation.

A Comprehensive Procedure to Evaluate the In Vivo Performance of Cancer Nanomedicines

The poor understanding of the in vivo performance of nanomedicines stymies their clinical translation. Procedures to evaluate the in vivo behavior of cancer nanomedicines at systemic, tissue, single-cell, and subcellular levels in tumor-bearing immunocompetent mice are described here. This approach may help researchers to identify promising cancer nanomedicines for clinical translation.

Inspired by the success of previous cancer nanomedicines in the clinic, researchers have generated a large number of novel formulations in the past ...

Cancer Research

In Vivo Tracking of Human Adipose-derived Mesenchymal Stem Cells in a Rat Knee Osteoarthritis Model with Fluorescent Lipophilic Membrane Dye

In order to support the clinical application of human adipose-derived mesenchymal stem cell (haMSC) therapy for knee osteoarthritis (KOA), we examined the efficacy of cell persistence and biodistribution of haMSCs in animal models. We demonstrated a method to label the cell membrane of haMSCs with lipophilic fluorescent dye. Subsequently, intra-articular injection of the labeled cells in rats with surgically induced KOA was monitored dynamically by an in vivo imaging system. We employed the lipophilic carbocyanines DiD (DilC18 (5)), a far-red fluorescent Dil (dialkylcarbocyanines) analog, which utilized a red laser to avoid excitation of the natural green autofluorescence from surrounding tissues. Furthermore, the red-shifted emission spectra of DiD allowed deep-tissue imaging in live animals and the labeling procedure caused no cytotoxic effects or functional damage to haMSCs. This approach has been shown to be an efficient tracking method for haMSCs in a rat KOA model. The application of this method could also be used to determine the optimal administration route and dosage of MSCs from other sources in pre-clinical studies.

In Vivo Tracking of Human Adipose-derived Mesenchymal Stem Cells in a Rat Knee Osteoarthritis Model with Fluorescent Lipophilic Membrane Dye

This protocol describes an efficient way to monitor the cell persistence and biodistribution of human adipose-derived mesenchymal stem cells (haMSCs) by far-red fluorescence labeling in a rat knee osteoarthritis (KOA) model via intra-articular (IA) injection.

In order to support the clinical application of human adipose-derived mesenchymal stem cell (haMSC) therapy for knee osteoarthritis (KOA), we examined the ...

Developmental Biology

Fluorescence-quenching of a Liposomal-encapsulated Near-infrared Fluorophore as a Tool for In Vivo Optical Imaging

Optical imaging offers a wide range of diagnostic modalities and has attracted a lot of interest as a tool for biomedical imaging. Despite the enormous number of imaging techniques currently available and the progress in instrumentation, there is still a need for highly sensitive probes that are suitable for in vivo imaging. One typical problem of available preclinical fluorescent probes is their rapid clearance in vivo, which reduces their imaging sensitivity. To circumvent rapid clearance, increase number of dye molecules at the target site, and thereby reduce background autofluorescence, encapsulation of the near-infrared fluorescent dye, DY-676-COOH in liposomes and verification of its potential for in vivo imaging of inflammation was done. DY-676 is known for its ability to self-quench at high concentrations. We first determined the concentration suitable for self-quenching, and then encapsulated this quenching concentration into the aqueous interior of PEGylated liposomes. To substantiate the quenching and activation potential of the liposomes we use a harsh freezing method which leads to damage of liposomal membranes without affecting the encapsulated dye. The liposomes characterized by a high level of fluorescence quenching were termed Lip-Q. We show by experiments with different cell lines that uptake of Lip-Q is predominantly by phagocytosis which in turn enabled the characterization of its potential as a tool for in vivo imaging of inflammation in mice models. Furthermore, we use a zymosan-induced edema model in mice to substantiate the potential of Lip-Q in optical imaging of inflammation in vivo. Considering possible uptake due to inflammation-induced enhanced permeability and retention (EPR) effect, an always-on liposome formulation with low, non-quenched concentration of DY-676-COOH (termed Lip-dQ) and the free DY-676-COOH were compared with Lip-Q in animal trials.

Fluorescence-quenching of a Liposomal-encapsulated Near-infrared Fluorophore as a Tool for In Vivo Optical Imaging

The use of fluorophores for in vivo imaging can be greatly limited by opsonization, rapid clearance, low detection sensitivity and cytotoxic effects on the host. Encapsulation of fluorophores in liposomes by film hydration and extrusion leads to fluorescence quenching and protection which enables in vivo imaging with high detection sensitivity.

Optical imaging offers a wide range of diagnostic modalities and has attracted a lot of interest as a tool for biomedical imaging. Despite the enormous ...

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 ...

Spatial Measurements of Perfusion, Interstitial Fluid Pressure and Liposomes Accumulation in Solid Tumors

The heterogeneous intra-tumoral accumulation of liposomes is a critical determinant of their efficacy. Both the chaotic tumor microcirculation and elevated IFP are linked to the heterogeneous intra-tumoral distribution of nanotechnology-based drug delivery systems such as liposomes. In the present study, the relationship between tumor microcirculation, elevated IFP, and accumulation of nanoparticles was investigated through in vivo experimentation. This was accomplished by evaluation of the tumor microcirculation using dynamic contrast enhanced computed tomography (DCE-CT) and measurement of tumor IFP using a novel image-guided robotic needle placement system connected to the micro-CT scanner. The intra-tumoral accumulation of liposomes was determined by CT image-based assessment of a nanoparticle liposomal formulation that stably encapsulate the contrast agent iohexol (CT-liposomes). CT imaging allowed for co-localization of the spatial distribution of tumor hemodynamics, IFP and CT-liposome accumulation in an individual subcutaneous xenograft mouse model of breast cancer. Measurements led to the discovery that perfusion and plasma volume fraction are strong mediators of the intra-tumoral distribution of liposomes. Furthermore, the results suggest that IFP plays an indirect role in mediating liposome distribution through modulating blood flow.

The heterogeneous intra-tumoral accumulation of liposomes has been linked to an abnormal tumor microenvironment. Herein methods are presented to measure tumor microcirculation by perfusion imaging and elevated interstitial fluid pressure (IFP) using an image-guided robotic system. Measurements are compared to the intra-tumoral accumulation of liposomes, determined using volumetric micro-CT imaging.

The heterogeneous intra-tumoral accumulation of liposomes is a critical determinant of their efficacy. Both the chaotic tumor microcirculation and ...

Medicine

A Quantitative Fluorescence Microscopy-based Single Liposome Assay for Detecting the Compositional Inhomogeneity Between Individual Liposomes

Most research employing liposomes as membrane model systems or drug delivery carriers relies on bulk read-out techniques and thus intrinsically assumes all liposomes of the ensemble to be identical. However, new experimental platforms able to observe liposomes at the single-particle level have made it possible to perform highly sophisticated and quantitative studies on protein-membrane interactions or drug carrier properties on individual liposomes, thus avoiding errors from ensemble averaging. Here we present a protocol for preparing, detecting, and analyzing single liposomes using a fluorescence-based microscopy assay, facilitating such single-particle measurements. The setup allows for imaging individual liposomes in a massive parallel manner and is employed to reveal intra-sample size and compositional inhomogeneities. Additionally, the protocol describes the advantages of studying liposomes at the single liposome level, the limitations of the assay, and the important features to be considered when modifying it to study other research questions.

This protocol describes the fabrication of liposomes and how these can be immobilized on a surface and imaged individually in a massive parallel manner using fluorescence microscopy. This allows for the quantification of the size and compositional inhomogeneity between single liposomes of the population.

Most research employing liposomes as membrane model systems or drug delivery carriers relies on bulk read-out techniques and thus intrinsically assumes ...

Using In Vitro Live-cell Imaging to Explore Chemotherapeutics Delivered by Lipid-based Nanoparticles

Conventional imaging techniques can provide detailed information about cellular processes. However, this information is based on static images in an otherwise dynamic system, and successive phases are easily overlooked or misinterpreted. Live-cell imaging and time-lapse microscopy, in which living cells can be followed for hours or even days in a more or less continuous fashion, are therefore very informative. The protocol described here allows for the investigation of the fate of chemotherapeutic nanoparticles after the delivery of doxorubicin (dox) in living cells. Dox is an intercalating agent that must be released from its nanocarrier to become biologically active. In spite of its clinical registration for more than two decades, its uptake, breakdown, and drug release are still not fully understood. This article explores the hypothesis that lipid-based nanoparticles are taken up by the tumor cells and are slowly degraded. Released dox is then translocated to the nucleus. To prevent fixation artifacts, live-cell imaging and time-lapse microscopy, described in this experimental procedure, can be applied.

Using In Vitro Live-cell Imaging to Explore Chemotherapeutics Delivered by Lipid-based Nanoparticles

Live-cell imaging is a powerful tool to visualize dynamic processes. The examination of fixed cells provides only static pictures, which can lead to misinterpretation and confusion about the process. This work presents a method to study uptake, drug release, and intracellular localization of liposomal nanoparticles in living cells.

Conventional imaging techniques can provide detailed information about cellular processes. However, this information is based on static images in an ...

Spatio-Temporal In Vivo Imaging of Ocular Drug Delivery Systems using Fiberoptic Confocal Laser Microendoscopy

Subconjunctival injection is an attractive route to administer ocular drugs due to easy trans-scleral access that bypasses anterior ocular barriers, such as the cornea and conjunctiva. While therapeutic effects and pharmacokinetics of the drugs upon subconjunctival injection have been described in some studies, very few assess the ocular distribution of drugs or drug delivery systems (DDS). The latter is critical for the optimization of intraocular DDS design and drug bioavailability to achieve the desired ocular localization and duration of action (e.g., acute versus. prolonged). This study establishes the use of fiberoptic confocal laser microendoscopy (CLM) to qualitatively study the ocular distribution of fluorescent liposomes in real-time in live mice after sub-conjunctival injection. Being designed for in vivo visual inspection of tissues at the microscopic level, this is also the first full description of the CLM imaging method to study spatio-temporal distribution of injectables in the eye after subconjunctival injection.

Spatio-Temporal In Vivo Imaging of Ocular Drug Delivery Systems using Fiberoptic Confocal Laser Microendoscopy

We present a protocol for the use of fiberoptic confocal laser microendoscopy (CLM) to non-invasively study the spatio-temporal distribution of liposomes in the eye after subconjunctival injection.

Subconjunctival injection is an attractive route to administer ocular drugs due to easy trans-scleral access that bypasses anterior ocular barriers, such ...

The Use of Ex Vivo Whole-organ Imaging and Quantitative Tissue Histology to Determine the Bio-distribution of Fluorescently Labeled Molecules

Fluorescent labeling is a well-established process for examining the fate of labeled molecules under a variety of experimental conditions both in vitro and in vivo. Fluorescent probes are particularly useful in determining the bio-distribution of administered large molecules, where the addition of a small-molecule fluorescent label is unlikely to affect the kinetics or bio-distribution of the compound. A variety of methods exist to examine bio-distribution that vary significantly in the amount of effort required and whether the resulting measurements are fully quantitative, but using multiple methods in conjunction can provide a rapid and effective system for analyzing bio-distributions.
Ex vivo whole-organ imaging is a method that can be used to quickly compare the relative concentrations of fluorescent molecules within tissues and between multiple types of tissues or treatment groups. Using an imaging platform designed for live-animal or whole-organ imaging, fluorescence within intact tissues can be determined without further processing, saving time and labor while providing an accurate picture of the overall bio-distribution. This process is ideal in experiments attempting to determine the tissue specificity of a compound or for the comparison of multiple different compounds. Quantitative tissue histology on the other hand requires extensive further processing of tissues in order to create a quantitative measure of the labeled compounds. To accurately assess bio-distribution, all tissues of interest must be sliced, scanned, and analyzed relative to standard curves in order to make comparisons between tissues or groups. Quantitative tissue histology is the gold standard for determining absolute compound concentrations within tissues.
Here, we describe how both methods can be used together effectively to assess the ability of different administration methods and compound modifications to target and deliver fluorescently labeled molecules to the central nervous system1.

The Use of Ex Vivo Whole-organ Imaging and Quantitative Tissue Histology to Determine the Bio-distribution of Fluorescently Labeled Molecules

Ex vivo whole-organ imaging is a rapid method for determining the relative concentrations of fluorescently labeled compounds within and between tissues or treatment groups. Conversely, quantitative fluorescence histology, while more labor intensive, allows for the quantification of the absolute tissue levels of labeled molecules.

Fluorescent labeling is a well-established process for examining the fate of labeled molecules under a variety of experimental conditions both in vitro ...

Biology

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.

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 ...

Preparation, Administration, and Assessment of In Vivo Tissue-Specific Cellular Uptake of Fluorescent Dye-Labeled Liposomes

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