We thank and acknowledge Dr. Peter Nelson and Sonya Anderson at the UK-ADC Brain Tissue Bank for providing all human brain tissue samples (NIH grant number: P30 AG028383 from the National Institute on Aging). We thank Matt Hazzard and Tom Dolan, Information Technology Services, Academic Technology and Faculty Engagement, University of Kentucky for graphical assistance. This project was supported by grant number 1R01NS079507 from the National Institute of Neurological Disorders and Stroke (to B.B.) and by grant number 1R01AG039621 from the National Institute on Aging (to A.M.S.H.). The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institute of Neurological Disorders and Stroke or the National Institute on Aging. The authors declare no competing financial interests.

The information below is based on current safety and regulatory standards at the University of Kentucky, Lexington, KY, USA. As a safety precaution, refer to the institution&#39;s biological safety program and the most current regulations and recommendations before working with human tissue.
CAUTION: Human tissue can be a source of blood-borne pathogens, including human immunodeficiency virus (HIV), hepatitis B virus (HBV), hepatitis C virus (HCV), and others. Working with human tissue poses the risk of infection from blood-borne pathogens. Therefore, certain regulatory and safety considerations are imperative when working with human tissue to protect laboratory personnel. Working with human tissue in the US requires a biosafety level 2 laboratory as well as safety precautions and training in accordance with NIH Section IV-B-7, OSHA Act of 1970 Clause 5(a)(1) and the user&#39;s institutional biological safety program. In general, Institutional Biosafety Committee and/or Institutional Review Board approval must be obtained prior to conducting any research involving human materials (tissue, body fluids). The training is required for all personnel working with human materials and includes basic laboratory safety training, e.g., Chemical Hygiene and Laboratory Safety, as well as specific training on biological safety, hazardous waste, and human blood-borne pathogens. All personnel working with human materials are highly recommended to obtain Hepatitis B vaccinations, prior to working with human materials. Personnel are required to wear specific personal protective equipment while working with human materials, e.g., a cuffed lab coat and a face shield, and wear gloves at all times. All work is performed in a biosafety cabinet (class 2). All equipment that comes in contact with human materials and any waste from human materials is handled appropriately to prevent contamination and/or infection of personnel. All equipment and surfaces are cleaned with 10% bleach and 75% ethanol following each procedure involving human materials. A spill with human materials must be immediately cleaned up. Glassware is autoclaved after each use. Waste, including unfixed human tissue, is collected in a labeled biohazard waste bag and autoclaved. Sharps are collected in a puncture- and leak-proof container labeled as biohazardous. All waste from human materials is disposed according to the institution&#39;s biological safety regulations.
NOTE: Our laboratory obtains fresh frontal cortex samples from deceased individuals through the Sanders-Brown Center on Aging (IRB #B15-2602-M). Inclusion criteria are: enrollment in the UK-ADC longitudinal autopsy cohort study and a Post-Mortem Interval (PMI) &#8804;4 h64. Autopsies follow a standard protocol and all procedures conform to NIH Biospecimen Best Practice Guidelines65. A short PMI of less than 4 h is of highest importance to ensure capillary viability after isolation. Both fresh and frozen tissue can be used. If freezing is necessary, freshly obtained human brain tissue should be shock-frozen in liquid nitrogen and stored at -80 &#176;C. Fresh or thawed tissue should be stored in isolation buffer (see below) and processed quickly. We find that 10 g of fresh human tissue yields about 100 mg of brain capillaries (wet weight).
1. Setup
<ol>
	<li>Buffer preparation 
	NOTE: The volume of buffer needed depends on the amount of tissue. All buffer volumes in the following protocol are based on 10 g of human brain cortex tissue.
	<ol>
		<li>L Isolation Buffer: Use 1.5 L of Dulbecco&#39;s phosphate-buffered saline (DPBS; 2.7 mM KCl, 1.47 mM KH2PO4, 136.9 mM NaCl, 8.1 mM Na2HPO4, 0.9 mM CaCl2, 0.49 mM MgCl2) and supplement with 5 mM D-glucose (1.35 g) and 1 mM sodium pyruvate (0.165 g). After adding the glucose and pyruvate, adjust to pH 7.4 with sodium hydroxide. Cool and store the buffer to 4 &#176;C prior to use.</li>
		<li>Bovine Serum Albumin (BSA): Add 10 g of BSA powder to 1 L of isolation buffer to a final BSA concentration of 1%. Stir slowly to avoid bubbles, adjust to pH 7.4, and store at 4 &#176;C overnight. Immediately prior to use, gently stir; avoid forming bubbles to avoid albumin denaturation.</li>
		<li>density gradient medium: Weigh 18 g of density gradient medium into a glass bottle and add a magnetic stir bar. Add 60 mL of isolation buffer and shake vigorously for 5 min until all powder is suspended. Store overnight at 4 &#176;C to allow the density gradient medium to dissolve. Stir for 10 min right before use.</li>
		<li>Store all buffers at 4 &#176;C; keep all tools and buffers on ice during the entire isolation procedure. Stir all buffers before use.</li>
	</ol>
	</li>
	<li>Experimental setup
	<ol>
		<li>Mount the pestle of the Potter-Elvehjem tissue grinder onto the electronic overhead stirrer. Place the Potter-Elvehjem tissue grinder and the Dounce homogenizer with pestle on ice under the hood. Prepare a 300 &#181;m filter mesh (5 x 5 cm2), fold it to a cone, and insert and attach it to a 50 mL Falcon tube with tape (Figure 1A).</li>
		<li>Place connecting rings and cell strain filters (pore size: 30 &#181;m) on 50 mL Falcon tubes. Prepare biohazardous waste bags. Place all required equipment in the biosafety cabinet (see Table of Materials).</li>
	</ol>
	</li>
</ol>
2. Brain Sample Preparation
NOTE: Figure 1A shows the workflow chart of the entire isolation procedure described below. Human brain tissue can stem from any part of the cortex and can be used fresh or frozen. Frozen brain tissue can be thawed at room temperature (no buffer; ~30 min for 10 g). To achieve comparable results, the brain tissue should be obtained from the same brain region for each experiment. This protocol is optimized for fresh (PMI &#60;4 h) human cerebral cortex that has not been frozen.
<ol>
	<li>Preparation of human brain tissue: Document the weight of the brain tissue. All numbers in the following protocol are appropriate for 10 g of fresh human brain tissue. Place the brain tissue in a 100 mm Petri dish. Carefully remove all the meninges with forceps. Use a scalpel to cut off the white matter.</li>
	<li>Mincing of the human brain tissue: Carefully cut up the brain tissue and mince it with a scalpel. Mince for about 5 min (2&#8211;3 mm pieces). Transfer the brain tissue to the Potter-Elvehjem tissue grinder. Add 30 mL of isolation buffer. 
	NOTE: The minced tissue pieces are difficult to see since the brain tissue turns into mush through the mincing process.</li>
</ol>
3. Homogenization
<ol>
	<li>Potter-Elvehjem tissue grinder (clearance: 150&#8211;230 &#181;m): Homogenize each sample with 100 strokes at a homogenizer speed of 50 rpm. Document the time every 25 strokes and the total time needed for 100 strokes. See Table 1 for a proposed homogenization protocol; the total time for homogenizing 10 g of human frontal cortex is about 22 min. Do not stir in air to prevent bubbles.</li>
	<li>Dounce homogenizer (clearance: 80&#8211;130 &#181;m): Transfer the homogenate to a Dounce homogenizer on ice. Homogenize the suspension with 20 strokes (~6 s/stroke, total of ~2 min). Avoid bubbles.</li>
</ol>
4. Centrifugation
<ol>
	<li>Distribute the brain homogenate equally into four 50 mL centrifugation tubes and document the total volume of the homogenate. Distribute 50 mL of density gradient buffer into the centrifugation tubes (12.5 mL per tube). Use 10 mL of isolation buffer to rinse the pestle and homogenizer, and distribute into the four centrifugation tubes (~2.5 mL per tube).</li>
	<li>Tightly close the centrifuge tubes with caps. Mix the homogenate, density gradient medium, and buffer by vigorously shaking the tubes. Centrifuge at 5,800 x g for 15 min at 4 &#176;C (fixed angle rotor); select a medium deceleration speed to keep the pellet attached to the tube. Discard the supernatant and resuspend each pellet in 2 mL of 1% BSA.</li>
</ol>
5. Filtration
NOTE: To separate the capillaries from red blood cells and other cell debris, several filtration steps are necessary.
<ol>
	<li>300 &#181;m mesh: After re-suspending the pellet, filter the suspension through the 300 &#181;m mesh. Capillaries are filtered through the mesh, whereas larger vessels and larger brain debris remain on the mesh. Carefully wash the mesh with up to 50 mL of 1% BSA. Discard the mesh. 
	NOTE: This filtration step clears the capillary suspension from any larger vessels or chunks of brain debris.</li>
	<li>30 &#181;M cell strain filter 
	NOTE: This filtration step separates capillaries from red blood cells and other brain debris.
	<ol>
		<li>Distribute the capillary filtrate from step 6.1 over the five 30 &#181;m cell strain filters (about 10 mL of capillary filtrate per cell strain filter). Capillaries are held back by this filter, whereas red blood cells, other single cells, and small brain debris pass through the filter and are collected in the filtrate.</li>
		<li>Wash each filter with 25 mL of 1% BSA. Afterwards, pour all filtrates over the sixth filter to increase the yield. Wash each filter with 50 mL of 1% BSA; keep the cell strain filters with containing the capillaries and discard the filtrate.</li>
	</ol>
	</li>
</ol>
6. Capillary Collection
<ol>
	<li>Turn the filters upside down and wash the capillaries with 50 mL of 1% BSA for each filter into 50 mL tubes. Gently apply pressure with the pipet tip of a 5 mL pipettor and move it across the filter to wash off the brain capillaries.</li>
	<li>Make sure to wash off all brain capillaries, especially from the rim of the filter. Avoid bubbles since this makes the filtration process more difficult and increases the chance of capillary loss.</li>
</ol>
7. Washing
<ol>
	<li>After collecting the capillaries, centrifuge all samples at 1,500 x g for 3 min at 4 &#176;C (swinging bucket rotor). Remove the supernatant and re-suspend the pellet in approximately 3 mL of isolation buffer. Combine all resuspended pellets from one sample in a 15 mL conical tube and fill it with isolation buffer. Centrifuge again at 1,500 x g for 3 min at 4 &#176;C and wash two more times.</li>
	<li>Document the capillary purity with a microscope (100X magnification) and camera (Figure 1B). 
	NOTE: The brain capillary yield from 10 g of human brain tissue is usually about 100 mg. The isolated brain capillaries can now be used for experiments, processed (e.g., lysate, membrane isolation), or be flash-frozen and stored at -80 &#176;C in cryotubes for a minimum of 6&#8211;12 months (avoid multiple freeze-thaw cycles).</li>
</ol>

The authors have nothing to disclose.

The present protocol describes the isolation of intact and viable human brain capillaries from fresh tissue. In this section, we discuss in detail the following: 1) modifications to the protocol, 2) troubleshooting of common errors, 3) limitations of the technique, 4) the significance of the model with respect to existing and alternative blood-brain barrier models, and 5) potential applications for isolated human brain capillaries.
The protocol described here is optimized for 10 g of fresh human frontal cortex tissue. However, it is relatively simple to modify this procedure for: 1) more or less than 10 g of tissue, 2) frozen brain tissue, or 3) brain tissue from a brain region other than the frontal cortex. First, with more or less than 10 g of brain tissue, the necessary volume of the buffers can simply be scaled up or down to the available amount of tissue. Thus, if only 5 g of brain tissue is available, the volume of the buffers should be reduced by half. Second, we describe a capillary isolation that used fresh brain tissue, but frozen tissue may be used if fresh tissue is unavailable38. Third, we used fresh brain tissue taken from the frontal cortex, but capillaries may be isolated from other cortical brain regions if there is enough available tissue. It is also possible to isolate capillaries from non-cortical brain regions (e.g., white matter), but these regions have a different cell composition and capillary density 66,67,68. Thus, using tissue from a different brain region would likely require the protocol to be adjusted (e.g., buffer volume, gradient medium density, centrifugation speed, and/or number of filtration steps).
The capillary isolation procedure, while not complex per se, is sensitive to small perturbations or alterations in the protocol. Modifications may result in a diminished capillary yield or reduced capillary viability. Table 2 outlines the most common errors and problems that are encountered during the isolation and lists tips to avoid these errors and solutions for troubleshooting if they occur. The most common problem associated with the procedure is a low capillary yield. The loss of capillaries is often the cumulative sum of small losses at each step and is due to small deviations across the procedure. A critical step in which a large amount of capillaries may be lost is the density centrifugation. An incorrect concentration in the buffer results in an incorrect density to separate the capillaries from cellular debris that reduces the volume of the capillary pellet. Figure 3 shows the consequences of too little or too much density gradient medium in the centrifugation step relative to brain homogenate. Adjusting to the correct the concentration may solve this problem. Note that the acceleration and deceleration speed of the centrifuge can also affect the formation of the brain capillary pellet. Capillaries may also be lost during steps 6&#8211;7 if part of the capillary material sticks to the pipette tips. This issue can be addressed by thoroughly rinsing each pipet tip before changing it. During step 7, washing off the capillaries from the cell strain filter may be incomplete and/or capillaries may stick to the rim of the filter. This can be avoided by checking the filter under the microscope followed by additional washing steps. Losing capillaries during each step of the isolation procedure can result in a negligible capillary pellet or not enough capillary material for further processing and experimentation.
Isolating brain capillaries from fresh human tissue represents a unique blood-brain barrier model that closely resembles the in vivo situation. However, several limitations of the technique exist. One challenge is the availability of fresh human tissue. As the optimal PMI is &#8804;4 h, brain tissue collected at a significantly longer PMI will not be fresh enough for some downstream applications. In some cases, it may be difficult to obtain tissue amounts that are large enough for multiple experimental groups, thereby restricting downstream applications. Thus, isolating fresh capillaries from rodent19, canine69, bovine42, or porcine70 brain tissue may be more suitable to model the blood-brain barrier. Factors that determine the variability of human brain tissue such as age, sex, ethnicity, disease state, medication history, brain region of sample, and PMI should be taken into consideration when interpreting and publishing data. On an experimental level it is important to note that isolated capillaries still include pericytes, but astrocytic endfeet are removed by the procedure44. It needs to be taken into consideration that the model presented here serves as an ex vivo model of the blood-brain barrier (i.e., capillary endothelial cells) but not as a model of the neurovascular unit.
Working with any human tissue always presents an inherent safety risk and researchers must take appropriate precautions during the isolation procedure to avoid infection. Specifically, in the U.S., work with human tissue requires designated laboratory space that is BSL 2-certified and includes a biosafety cabinet (class A2). In addition, staff must use personal protective equipment (i.e., lab coat, gloves, and face shield) and designated equipment for work with human tissue and implement biohazardous waste handling. Implementing these safety measures is time-consuming, cost-intensive, and increases the difficulty of the procedure, especially for inexperienced laboratory personnel.
The blood-brain barrier is highly conserved among organisms with a well-defined CNS71. Modeling the human blood-brain barrier is difficult because there is complex neurovascular coupling among the cells of the neurovascular unit. An adult human brain has been estimated to have in average about 86 billion neurons and it is thought that almost every neuron has its own capillary in the vicinity to ensure proper supply with oxygen and nutrients72,73. Capillary endothelial cells constitute the largest surface area of the blood-brain interface (12&#8211;18 m2&#160;for a healthy adult human). Tight junctions represent an impediment to a wide array of pharmacotherapeutics by blocking paracellular diffusion of solutes. In addition, numerous studies describe barrier dysfunction in neurodegenerative disorders, e.g., Alzheimer&#39;s disease74, stroke75, epilepsy38,76, multiple sclerosis77, and traumatic brain injury28,78. Thus, it is imperative to establish models that closely represent the human blood-brain barrier and allow for a better understanding of barrier function in health and disease.
Numerous in vitro cell culture models of the blood-brain barrier exist; for expert reviews on the subject see 41,49,51,69,79,80. Briefly, both freshly isolated brain capillary endothelial cells for primary culture and immortalized brain capillary endothelial cell lines are available. Primary cultures of cerebral microvessel endothelial cells are mostly used from mouse, rat, pig, and cow. However, primary cell cultures are labor-intensive since cells must be freshly isolated. Immortalized brain capillary endothelial cell lines are available from mouse, rat, and human and are less labor-intensive because they can be passaged for longer-term use. However, even immortalized cell lines have a limit on how often they can be passaged before losing their endothelial characteristics. Both primary cells as well as immortalized cells lines are often cultured on plates to model the brain capillary endothelium and measure the barrier permeability and drug transport across the cell monolayer, thereby mimicking blood-to-brain transport41,81,82. The culture media in these models may also be modified or supplemented with astrocytes, pericytes, or other physiologically relevant factors like cAMP41,83,84.
The advantage of immortalized cell lines is their relatively easy access and availability. While cultured endothelial cells can reach confluence, they lose endothelial cell properties as they grow side-by-side in a monolayer. For example, cultured hCMECs display reduced expression of transporters like P-gp, tight junction proteins, and display variable permeability to xenobiotics41. Figure 4 shows a Western blot for P-gp protein expression in hCMEC/D3 cells compared to freshly isolated human capillaries. Despite a 10-fold lower amount of total protein, the signal for P-gp is stronger in isolated human capillaries compared to hCMEC/D3 cells. This indicates that hCMEC/D3 cells lost a significant amount of P-gp protein expression in culture. Moreover, differences in culture media, environment, and equipment affect key measures of barrier integrity, i.e., TEER measurements in Transwell plate assays. Some of these issues can be overcome by utilizing the isolated brain capillary model that more closely represents the human blood-brain barrier in vivo.
Isolated brain capillaries have been used for a wide range of studies, including genomics, proteomics, functional proteomics, and cellomics studies (Figure 5). In addition, many techniques and methods exist to analyze isolated brain capillaries within each of these fields. Notably, the experimental techniques shown in Figure 5 can be used on brain capillaries isolated from a number of sources, including human, bovine, rodent, and porcine tissue, which may facilitate translational research. For example, Li et al.85 studied the genomics of the blood-brain barrier using suppression subtractive hybridization by purifying mRNA isolated from rat brain capillaries. Additionally, Ott et al.86 used RT-PCR and qRT-PCR to study the regulation of P-gp by PXR. Many proteomic studies utilize Western blotting3, Dot Blot analysis87, Simple Western assays3,38, ELISA88,89, immunoprecipitation3, and immunostaining3,90,91. To discern transporter trafficking in the brain endothelium, McCaffrey et al.92 used subcellular fractionation of isolated brain capillaries. Sa&#769;nchez del Pino et al.93used isolated bovine endothelial membrane vesicles to discern transporter location and transport direction across the blood-brain barrier. In other proteomic studies, researchers used liquid chromatography and tandem mass spectrometry to quantify transporter proteins94,95,96. Functional proteomic studies have utilized transporter and leakage assays2,38. Hartz et al.2 used zymography to determine enzyme activity in isolated brain capillaries. In addition, vast cellomic research using cell culture has generated numerous endothelial cell lines and models of the blood-brain barrier82,83,84. With these models, common assays used include migration assays97,98, viability and toxicity assays99, and angiogenesis assays98.
Isolated brain capillaries allow for accurate characterization of protein expression and activity and description of signaling pathways at the blood-brain barrier. This is due, in part, to the capillary content of the brain, which is only approximately 1% (v/v). Thus, using whole brain homogenate or brain slices as a substitute for purified capillary endothelial cells will most likely result in a poor signal-to-noise ratio24. In addition, after isolation, brain capillaries are viable for at least 6 h (unpublished data from mouse and rat), which allows studies to discern specific signaling pathways. It is recommended to include a control group from the same preparation.
Representative and translational models of the blood-brain barrier, such as the isolated capillary model discussed in this report, are needed to study barrier function in health and disease. Here we present a protocol to obtain isolated human brain capillaries at a good yield and high quality that can serve as an ex vivo model of the blood-brain barrier. Isolated capillaries retain their original structure and function, which allows using them for a number of post-isolation molecular, biochemical, and physiological assays. Care should be taken when handling human tissue samples, preferably in a BSL 2 or higher setting.

The blood-brain barrier is a tightly controlled interface between the blood and brain that determines what goes into and comes out of the brain. Anatomically, endothelial cells compose the blood-brain barrier and forms a complex, continuous capillary network. Physiologically, this capillary network supplies the brain with oxygen and nutrients while simultaneously disposing of carbon dioxide and metabolic waste products. Importantly, evidence supports that the changes to the barrier contribute to numerous pathologies, including Alzheimer&#39;s disease, epilepsy, and stroke1,2,3,4,5,6,7. Brain endothelial cells also serve as a barrier to treatment by blocking drug uptake into the brain, e.g., chemotherapy of glioblastoma multiforme following tumor resection8,9,10. In this regard, isolated human brain capillaries represent a unique ex vivo blood-brain barrier model that closely resembles barrier properties in vivo, which allows for the study of barrier function and dysfunction in health and disease. In this article, we provide a protocol to isolate brain capillaries from human brain at a consistently high capillary quality and yield to study the blood-brain barrier.
In 1969, Siakotos et al.11 were the first to report the isolation of brain capillaries from bovine and human brain tissue using density gradient centrifugation and glass bead column separation. Later, Goldstein et al.12 improved this method by adding multiple filtration steps to decrease the amount of tissue needed to study brain capillaries isolated from rats, while maintaining the metabolic activity of glucose transport. Since then, researchers optimized the capillary isolation procedure numerous times, improving the method and brain capillary model with each iteration13,14,15. For example, Pardridge et al.16 isolated bovine capillaries using enzymatic digestion rather than mechanical homogenization, and then subsequently passed a capillary suspension through a 210 &#181;m mesh filter and a glass bead column. These modifications improved the trypan blue exclusion stain of isolated brain capillaries, and thus, increased endothelial cell viability. In the early 1990s, Dallaire et al.17 isolated bovine and rat capillaries that were clear of neuronal contamination and maintained metabolic activity of &#947;-glutamyl transpeptidase (&#947;-GTase) and alkaline phosphatase. In 2000, Miller et al.18, used isolated rat and porcine brain capillaries in combination with confocal microscopy to show the accumulation of transport substrates into the lumen of capillaries. Subsequently, our laboratory has continued to optimize the brain capillary isolation procedure and we have established transport assays to determine P-glycoprotein (P-gp)19,20,21, breast cancer resistance protein (BCRP)22,23, and multi-drug resistance protein 2 (Mrp2)24 transport activity. In 2004, we published two reports where we used isolated rat brain capillaries to investigate various signaling pathways. In Hartz et al.21, we found that the peptide endothelin-1 rapidly and reversibly reduced P-gp transport function in brain capillaries by acting through the endothelin receptor B (ETB) receptor, nitric oxide synthase (NOS), and protein kinase C (PKC). In Bauer et al.19, we demonstrated expression of the nuclear receptor pregnane X receptor (PXR) and showed PXR-modulation of P-gp expression and transport function in brain capillaries. In experiments with transgenic humanized PXR mice, we expanded this line of research and showed in vivo tightening of the barrier by upregulating P-gp through hPXR activation25. In 2010, Hartz et al.26 used this approach to restore P-gp protein expression and transport activity in transgenic human amyloid precursor protein (hAPP) mice that overexpress hAPP. Moreover, restoring P-gp in hAPP mice significantly reduced amyloid beta (A&#946;)40and A&#946;42brain levels.
In addition to studying signaling pathways, isolated brain capillaries can be used to determine changes in capillary permeability which we refer to as capillary leakage. In particular, the Texas Red leakage assay is used to assess leakage of the fluorescent dye Texas Red from the capillary lumen over time and these data are then used to analyze leakage rates. Increased capillary leakage rates compared to those from control capillaries indicate changes in the physical integrity of the blood-brain barrier2. This is valuable because there are numerous disease states associated with barrier disruption, e.g., epilepsy, multiple sclerosis, Alzheimer&#39;s disease, and traumatic brain injury27,28,29,30. Other groups have also utilized isolated capillaries to discern signaling pathways that regulate protein expression and transport activity of proteins31,32,33,34,35,36,37. Finally, we have continued to optimize this method for the isolation of human brain capillaries and, recently, we showed increased P-gp expression at the human blood-brain barrier in patients with epilepsy compared to seizure-free control individuals38. Taken together, these developments demonstrate that isolated brain capillaries can serve as a versatile model to study barrier function.
Various in vivo, ex vivo, and in vitro blood-brain barrier models have been used in basic research and industrial drug screening, mainly with the goal of testing drug delivery to the brain39,40,41,42,43,44. In addition to isolated ex vivo brain capillaries, current blood-brain barrier models include in silico models, in vitro cell culture of isolated brain capillary endothelial cells or immortalized cell lines from various species, in vitro culture of human pluripotent stem cells (hPSC) that differentiate into brain capillary endothelial cells, and microfluidic models on a chip.
In silico models are most commonly used in drug development for selecting drug candidates based on predicted absorption, distribution, metabolism, and excretion (ADME) properties. Methods such as quantitative structure-property relationship (QSPR) models and quantitative structure-activity relationship (QSAR) models are popular methods used in high-throughput screening of libraries to predict brain penetration of drug candidates45,46. These models are useful to screen molecules for barrier penetration properties.
Betz et al.47 established monolayers of cultured brain capillary endothelial cells as an in vitro blood-brain barrier model system. In vitro cell culture models using fresh tissue or immortalized endothelial cell lines such as human cerebral microvessel endothelial cells (hCMECs) can be another high-throughput screening tool for brain penetration or mechanistic studies. However, brain capillary endothelial cell culture models lack the physiologic shear stress of blood flow inside the capillary lumen, are limited in overall biologic complexity, and undergo changes in expression and localization of important barrier components such as tight junction proteins, surface receptors, transporters, enzymes, and ion channels48,49,50. Conversely, endothelial monolayers derived from hPSCs, have low sucrose permeability compared to hCMEC/D3 cultures and contain polarized expression of some blood-brain barrier transporters, adhesion molecules, and tight junctions51,52. However, these cells are also subject to changing properties in the culture, and the system must be validated for its recapitulation of in vivo barrier properties52.
Newer trends in blood-brain barrier research include utilizing 3D tissue culture systems to create artificial capillaries, using the organ-on-chip technology to generate microfluidic devices, or utilizing the hollow fiber technology53,54,55. Artificial capillaries, however, have significantly larger diameters (100&#8211;200 &#181;m) than brain capillaries (3&#8211;7 &#181;m). Hence, the shear forces in vitro do not fully resemble the in vivo situation. This is addressed in &#34;blood-brain-barrier-on-a-chip&#34; microfluidic devices, where artificial membranes form &#34;blood&#34; and &#34;brain&#34; compartments and fluids are pumped through these devices generating microfluidic shear forces. Similarly, co-cultures of endothelial cells in various combinations with astrocytes and vascular smooth muscle cells have also been used with the hollow fiber technology to recreate rheological parameters present under in vivo conditions56,57,58. However, it is unclear how well this model reflects other properties of the blood-brain barrier such as transport, metabolism, signaling, and others. These artificial capillary and chip models are suitable for high-throughput screening of drugs, but the cells used to generate these models are also subject to change during culture.
Frozen and fixed brain slices or primary brain capillary endothelial cell cultures are additional models that can be used tostudy the human microvasculature5,59,60,61. For example, immunohistochemistry of fixed brain tissue is used to determine protein localization and expression in healthy compared to diseased tissue.
In addition to tissue slices and the in vitro models described above, freshly isolated brain capillaries can be utilized to study blood-brain barrier function. Limitations of this isolated capillary model include the difficulty to obtain fresh human brain tissue, absence of astrocytes and neurons, and a relatively time-consuming isolation process. An advantage of the isolated brain capillary model is that this model closely resembles the in vivo situation and, therefore, can be used to characterize barrier function and dysfunction. Importantly, it can also be used to discern signaling mechanisms using a multitude of assays and molecular techniques3,19,62,63.
Our laboratory has access to both fresh and frozen human brain tissue through the Sanders-Brown Center on Aging (IRB #B15-2602-M)64. In this context, autopsies follow a standard protocol, brains are obtained in &#60;4 h, and all procedures conform to NIH Biospecimen Best Practice Guidelines65. Given this unique access to human brain tissue, we established and optimized a protocol to isolate brain capillaries from human brain tissue that results in a high yield of intact, viable human brain capillaries. Two common endpoints of interest are to determine the protein expression and activity. In this regard, we and others have established various assays that can be used with isolated brain capillaries to study protein expression and activity levels. These assays include Western blotting, Simple Western assay, enzyme-linked immunosorbent assay (ELISA), reverse transcription polymerase chain reaction (RT-PCR), quantitative polymerase chain reaction (qPCR), zymography, transport activity assays, and capillary leakage assays. These assays allow researchers to study changes in barrier function in human pathologic conditions, determine pathways that govern protein expression and activity, and identify pharmacologic targets for the treatment of blood-brain barrier associated diseases.
Taken together, freshly isolated brain capillaries can serve as a robust and reproducible model of the blood-brain barrier. Especially, this model can be combined with many different assays to determine a wide array of endpoints to study barrier function.

<table>
	<tbody>
		<tr>
			<td>Personal Protective Equipment (PPE)</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Diamond Grip Plus Latex Gloves, Microflex Medium</td>
			<td>VWR, Radnor, PA, USA</td>
			<td>32916-636</td>
			<td>PPE</td>
		</tr>
		<tr>
			<td>Disposable Protective Labcoats</td>
			<td>VWR, Radnor, PA, USA</td>
			<td>470146-214</td>
			<td>PPE; due to the nature of the human source material, the use of a disposable lab coat is recommended</td>
		</tr>
		<tr>
			<td>Face Shield, disposable</td>
			<td>Thermo Fisher Scientific, Pittsburgh, PA, USA</td>
			<td>19460102</td>
			<td>PPE; due to the nature of the human source material, the use of a disposable face shield is recommended</td>
		</tr>
		<tr>
			<td>Safety Materials</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Clavies High-Temperature Autoclave Bags 8X12</td>
			<td>Thermo Fisher Scientific, Pittsburgh, PA, USA</td>
			<td>01-815-6</td>
			<td></td>
		</tr>
		<tr>
			<td>Versi Dry Bench Paper 18&#34; x 20&#34;</td>
			<td>Thermo Fisher Scientific, Pittsburgh, PA, USA</td>
			<td>14-206-32</td>
			<td>to cover working areas</td>
		</tr>
		<tr>
			<td>VWR Sharps Container Systems</td>
			<td>Thermo Fisher Scientific, Pittsburgh, PA, USA</td>
			<td>75800-272</td>
			<td>for used scalpels</td>
		</tr>
		<tr>
			<td>Bleach 8.2% Clorox Germicidal 64 oz</td>
			<td>UK Supply Center, Lexington, KY, USA</td>
			<td>323775</td>
			<td></td>
		</tr>
		<tr>
			<td>Equipment</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>4&#176;C Refrigerator</td>
			<td>Thermo Fisher Scientific, Pittsburgh, PA, USA</td>
			<td>13-986-148</td>
			<td></td>
		</tr>
		<tr>
			<td>Accume BASIC AB15 pH Meter</td>
			<td>Thermo Fisher Scientific, Pittsburgh, PA, USA</td>
			<td>AB15</td>
			<td></td>
		</tr>
		<tr>
			<td>Heidolph RZR 2102 Control</td>
			<td>Heidolph, Elk Grove Village, IL, USA</td>
			<td>501-21024-01-3</td>
			<td></td>
		</tr>
		<tr>
			<td>Sorvall LEGEND XTR Centrifuge</td>
			<td>Thermo Fisher Scientific, Pittsburgh, PA, USA</td>
			<td>75004521</td>
			<td></td>
		</tr>
		<tr>
			<td>Leica L2 Dissecting Microscope</td>
			<td>Leica Microsystems Inc, Buffalo Grove IL, USA</td>
			<td></td>
			<td>used to remove meninges</td>
		</tr>
		<tr>
			<td>POLYTRON PT2500 Homogenizer</td>
			<td>Kinematica AG, Luzern, Switzerland</td>
			<td>9158168</td>
			<td></td>
		</tr>
		<tr>
			<td>Scale P-403</td>
			<td>Denver Instrument, Bohemia, NY, USA</td>
			<td>0191392</td>
			<td></td>
		</tr>
		<tr>
			<td>Standard mini Stir</td>
			<td>Thermo Fisher Scientific, Pittsburgh, PA, USA</td>
			<td>1151050</td>
			<td></td>
		</tr>
		<tr>
			<td>Thermo-Flasks Liquid Nitrogen Dewar</td>
			<td>Thermal Scientific, Mansfiled, TX, USA</td>
			<td>11-670-4C</td>
			<td>used to freeze the tissue?</td>
		</tr>
		<tr>
			<td>Voyager Pro Analytical Balance</td>
			<td>OHAUS, Parsippany, NJ, USA</td>
			<td>VP214CN</td>
			<td></td>
		</tr>
		<tr>
			<td>ZEISS Axiovert Microcope</td>
			<td>Carl Zeiss, Inc Thornwood, NY, USA</td>
			<td></td>
			<td>used to check isolated capillaries</td>
		</tr>
		<tr>
			<td>Tools and Glassware</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Finnpipette II Pipette 1-5mL</td>
			<td>Thermo Fisher Scientific, Pittsburgh, PA, USA</td>
			<td>21377823T1</td>
			<td>wash capillaries off filter</td>
		</tr>
		<tr>
			<td>Finnpipette II Pipette 100-1000 &#181;L</td>
			<td>Thermo Fisher Scientific, Pittsburgh, PA, USA</td>
			<td>21377821T1</td>
			<td>resuspend pellet in BSA</td>
		</tr>
		<tr>
			<td>Pipet Boy</td>
			<td>Integra, Hudson, NH, USA</td>
			<td>739658</td>
			<td></td>
		</tr>
		<tr>
			<td>50mL Falcon tubes 25/rack - 500/cs</td>
			<td>VWR, Radnor, PA, USA</td>
			<td>21008-951</td>
			<td></td>
		</tr>
		<tr>
			<td>EISCO Scalpel Blades</td>
			<td>Thermo Fisher Scientific, Pittsburgh, PA, USA</td>
			<td>S95938C</td>
			<td>to mince brain tissue</td>
		</tr>
		<tr>
			<td>PARAFILM</td>
			<td>VWR, Radnor, PA, USA</td>
			<td>52858-000</td>
			<td>to cover beaker and volumetric flask</td>
		</tr>
		<tr>
			<td>Thermo Scientific Finntip Pipet Tips 5 ml</td>
			<td>Thermo Fisher Scientific, Pittsburgh, PA, USA</td>
			<td>21-377-304</td>
			<td>to wash capillaries off filter</td>
		</tr>
		<tr>
			<td>60 ml syringe with Luer-Lok</td>
			<td>Thermo Fisher Scientific, Pittsburgh, PA, USA</td>
			<td>BD309653</td>
			<td>used with connector ring to filter capillaries</td>
		</tr>
		<tr>
			<td>Scalpel Handle #4</td>
			<td>Fine Science Tools, Foster City, CA, USA</td>
			<td>10060-13</td>
			<td>used for mincing</td>
		</tr>
		<tr>
			<td>Dumont Forceps #5</td>
			<td>Fine Science Tools, Foster City, CA, USA</td>
			<td>11251-10</td>
			<td>used to remove meninges</td>
		</tr>
		<tr>
			<td>Potter-Elvehjem Tissue Grinder</td>
			<td>Thomas Scientific, Swedesboro, NJ, USA</td>
			<td>3431E25</td>
			<td>50 ml volume, clearance: 150-230 &#956;m</td>
		</tr>
		<tr>
			<td>Dounce Homogenizer</td>
			<td>VWR, Radnor PA USA</td>
			<td>62400-642</td>
			<td>15 ml volume, clearance: 80-130 &#956;m</td>
		</tr>
		<tr>
			<td>Spectra/Mesh Woven Filters (300 &#181;m)</td>
			<td>Spectrum Laboratories, Rancho Dominguez, CA, USA</td>
			<td>146424</td>
			<td>Used to filter capillary suspension to remove any meninges that may be left</td>
		</tr>
		<tr>
			<td>pluriStrainers (pore size: 30 &#181;m)</td>
			<td>pluriSelect Life Science, Leipzig, Germany</td>
			<td>43-50030-03</td>
			<td></td>
		</tr>
		<tr>
			<td>Connector Ring</td>
			<td>pluriSelect Life Science, Leipzig, Germany</td>
			<td>41-50000-03</td>
			<td>reuse multiple time</td>
		</tr>
		<tr>
			<td>1 l Volumetric Flask</td>
			<td></td>
			<td></td>
			<td>for preparation of Isolation Buffer</td>
		</tr>
		<tr>
			<td>1 l Beaker</td>
			<td></td>
			<td></td>
			<td>for preparation of 1% BSA</td>
		</tr>
		<tr>
			<td>Stir Bar</td>
			<td></td>
			<td></td>
			<td>for preparation of 1% BSA and Ficoll&#174;</td>
		</tr>
		<tr>
			<td>Schott Bottle (60 ml)</td>
			<td></td>
			<td></td>
			<td>for preparation of Ficoll&#174;</td>
		</tr>
		<tr>
			<td>Ice Bucket</td>
			<td></td>
			<td></td>
			<td>to keep everything cold</td>
		</tr>
		<tr>
			<td>100 mm Petri Dish</td>
			<td></td>
			<td></td>
			<td>for mincing of brain tissue</td>
		</tr>
		<tr>
			<td>Tissue Culture Cell Scraper</td>
			<td>VWR, Radnor, PA, USA</td>
			<td>89260-222</td>
			<td>to remove supernatant after centrifugation</td>
		</tr>
		<tr>
			<td>Chemicals</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>BSA Fraction V, A-9647</td>
			<td>Sigma-Aldrich, St. Louis, MO, USA</td>
			<td>A9647-500g</td>
			<td>prepare in DPBS with Ca2+ &#38; Mg2+ the day before. Avoid bubbles during preparation. Store in the refrigerator. Slowly stir for 10 min before use.</td>
		</tr>
		<tr>
			<td>DPBS with Ca2+ &#38; Mg2+</td>
			<td>Hyclone</td>
			<td>SH30264.FS</td>
			<td>DPBS - part of the Isolation Buffer</td>
		</tr>
		<tr>
			<td>Ficoll PM400</td>
			<td>Sigma-Aldrich, St. Louis, MO, USA</td>
			<td>F4375</td>
			<td>Exact measurement is important here. Weigh out in bottle with stir bar. Shake vigurously after adding DPBS. Keep in the fridge O/N. It will be clear in the morning. Stir gently for 10-15 min before use. Keep on ice until use.</td>
		</tr>
		<tr>
			<td>Glucose (D-(+) Dextrose)</td>
			<td>Sigma-Aldrich, St. Louis, MO, USA</td>
			<td>G7528</td>
			<td>Glucose (D-(+) Dextrose) Concentration: 5 mM</td>
		</tr>
		<tr>
			<td>Sodium Hydroxide Standard Solution</td>
			<td>Sigma-Aldrich, St. Louis, MO, USA</td>
			<td>71474</td>
			<td>to adjust pH of the DPBS</td>
		</tr>
		<tr>
			<td>Sodium Pyruvate</td>
			<td>Sigma-Aldrich, St. Louis, MO, USA</td>
			<td>P2256</td>
			<td>Concentration: 1 mM</td>
		</tr>
	</tbody>
</table>

isolation of cerebral capillaries from fresh human brain tissue

Understanding blood-brain barrier function under physiological and pathophysiological conditions is critical for the development of new therapeutic strategies that hold the promise to enhance brain drug delivery, improve brain protection, and treat brain disorders. However, studying the human blood-brain barrier function is challenging. Thus, there is a critical need for appropriate models. In this regard, brain capillaries isolated from human brain tissue represent a unique tool to study barrier function as close to the human in vivo situation as possible. Here, we describe an optimized protocol to isolate capillaries from human brain tissue at a high yield and with consistent quality and purity. Capillaries are isolated from fresh human brain tissue using mechanical homogenization, density-gradient centrifugation, and filtration. After the isolation, the human brain capillaries can be used for various applications including leakage assays, live cell imaging, and immune-based assays to study protein expression and function, enzyme activity, or intracellular signaling. Isolated human brain capillaries are a unique model to elucidate the regulation of the human blood-brain barrier function. This model can provide insights into central nervous system (CNS) pathogenesis, which will help the development of therapeutic strategies for treating CNS disorders.

The isolations from human brain tissue yield a suspension enriched in human brain capillaries (Figure 1B) with small amounts of larger vessels, red blood cells, other single cells, and some cell debris. Some capillaries are branched, and, in some, red blood cells are entrapped in the capillary lumens. The typical capillary has a 3&#8211;7 &#181;m diameter and is approximately 100-200 &#181;m long with open lumens; most capillary ends are collapsed. Using confocal microscopy, isolated human brain capillaries reveal a tubular, intact structure and morphology. Figure 2A shows a representative transmitted light image of a human brain capillary with an attached pericyte and a red blood cell in the lumen. All of the findings regarding diameter, size, and morphology are in accordance with previous reports on the structure of isolated brain capillaries12,17,18. The isolated human brain capillary in Figure 2B was immunostained for P-gp (green) using C219 as the primary antibody (1 &#181;g/mL); nuclei were counterstained with DAPI (1 &#181;g/mL).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/57346/57346fig1.jpg" /> 
Figure 1: Flowchart for capillary isolation. (A) The pictogram illustrates major steps of the procedure to isolate brain capillaries from fresh human tissue. (B) The picture shows isolated human brain capillaries under a light microscope directly after isolation (100X magnification). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/57346/57346fig1large.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/57346/57346fig2.jpg" /> 
Figure 2: Isolated human brain capillary. (A) A transmitted light image of an isolated human brain capillary. (B) The confocal microscope image shows an isolated human brain capillary immunostained for P-gp (green; C219 1 &#181;g/mL); nuclei were counterstained with DAPI (blue; 1 &#181;g/mL). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/57346/57346fig2large.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/57346/57346fig3.jpg" /> 
Figure 3: Troubleshooting density centrifugation. The pictogram shows the preparation after the density gradient centrifugation. It highlights the effects of too much and too little density gradient medium and how this affects the separation and the resulting capillary pellet. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/57346/57346fig3large.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/57346/57346fig4.jpg" /> 
Figure 4: P-gp protein expression in isolated human brain capillaries. The Western blot shows strong bands for P-gp (1 &#181;g/mL) in isolated human capillaries compared to hCMEC\D3 cells. &#946;-Actin was used as a loading control (1 &#181;g/&#181;L). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/57346/57346fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/57346/57346fig5.jpg" /> 
Figure 5: Applications for isolated human brain capillaries. An overview of the most common applications for isolated brain capillaries published in the literature. Isolated capillaries have been used for: 1) Genomics85,86, 2) Proteomics3,38,87,88,89,90,91,92,94,95,96, 3) Functional Proteomics2,38, and 4) Cellomics82,83,84,97,98,99. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/57346/57346fig5large.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>Strokes</td>
			<td>Time [min]</td>
		</tr>
		<tr>
			<td>1&#8211;25</td>
			<td>7&#8211;7.5</td>
		</tr>
		<tr>
			<td>26&#8211;50</td>
			<td>5&#8211;5.5</td>
		</tr>
		<tr>
			<td>51&#8211;75</td>
			<td>5&#8211;5.5</td>
		</tr>
		<tr>
			<td>76&#8211;100</td>
			<td>5&#8211;5.5</td>
		</tr>
		<tr>
			<td>Total Time:</td>
			<td>22&#8211;24 min</td>
		</tr>
	</tbody>
</table>
Table 1: Homogenization protocol. The homogenization protocol for the Potter-Elvehjem tissue grinder to homogenize 10 g of human frontal cortex at a homogenization speed of 50 rpm. Note that the first several strokes require additional time to homogenize the minced tissue. After this initial homogenization, each stroke is 12 s in duration (6 s for downward movement, 6 s for upward movement). Thus, after the initial homogenization, 5 strokes can be accomplished in 1 min, or 25 strokes in 5 min.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Problem</td>
			<td>Potential Cause</td>
			<td>Solution</td>
		</tr>
		<tr>
			<td rowspan="3">No Capillary Pellet</td>
			<td>1)&#160; Incorrect Ficoll concentration</td>
			<td>1)&#160; Adjust Ficoll concentration</td>
		</tr>
		<tr>
			<td>2)&#160; Incorrect centrifugation speed</td>
			<td>2)&#160; Adjust centrifugation speed</td>
		</tr>
		<tr>
			<td>3)&#160; Incorrect acceleration and/or deceleration speed</td>
			<td>3)&#160; Adjust acceleration and/or deceleration speed</td>
		</tr>
		<tr>
			<td rowspan="4">Low Capillary Yield</td>
			<td>1)&#160; Meninges blocking filtration steps</td>
			<td>1)&#160; Remove ALL meninges prior to filtration</td>
		</tr>
		<tr>
			<td>2)&#160; Too many capillaries lost during isolation procedure</td>
			<td>2)&#160; Calculate buffer concentration correctly, rinse pipette tips</td>
		</tr>
		<tr>
			<td>3)&#160; Washing capillaries off PluriStrainer filters was insufficient</td>
			<td>3)&#160; Turn filters over and carefully inspect for capillaries (use microscope)</td>
		</tr>
		<tr>
			<td>4)&#160; Excess bubbles during resuspensions</td>
			<td>4)&#160; &#160;Pipette slowly to avoid bubbles</td>
		</tr>
		<tr>
			<td rowspan="4">Non-Viable Capillaries</td>
			<td>1)&#160; Extended post-mortem interval</td>
			<td>1)&#160; Reduce interval if possible or use snap-frozen brains</td>
		</tr>
		<tr>
			<td>2)&#160; Using frozen tissue for the isolation</td>
			<td>2)&#160; Use fresh tissue</td>
		</tr>
		<tr>
			<td>3)&#160; Time of isolation procedure too long</td>
			<td>3)&#160; Optimize workflow</td>
		</tr>
		<tr>
			<td>4)&#160; Equipment/buffers were not kept on ice during the isolation procedure</td>
			<td>4)&#160; Keep equipment and buffers on ice during the isolation</td>
		</tr>
	</tbody>
</table>
Table 2: Troubleshooting of common problems. A list of the most common errors and problems that occur during the isolation procedure and how they can be resolved.

Watch this Scientific Journal Video about Isolation of Cerebral Capillaries from Fresh Human Brain Tissue at JoVE.com

Isolation of Cerebral Capillaries from Fresh Human Brain Tissue

Isolated brain capillaries from human brain tissue can be used as a preclinical model to study barrier function under physiological and pathophysiological conditions. Here, we present an optimized protocol to isolate brain capillaries from fresh human brain tissue.

Understanding blood-brain barrier function under physiological and pathophysiological conditions is critical for the development of new therapeutic ...

isolation-of-cerebral-capillaries-from-fresh-human-brain-tissue

Nanyang Technological University

Research

JoVE Journal

Neuroscience

12.5K Views.  University of Kentucky. Isolated brain capillaries from human brain tissue can be used as a preclinical model to study barrier function under physiological and pathophysiological conditions. Here, we present an optimized protocol to isolate brain capillaries from fresh human brain tissue.

Video: Isolation of Cerebral Capillaries from Fresh Human Brain Tissue

Double Fluorescence in situ Hybridization in Fresh Brain Sections

Here we describe a modified version of a double fluorescence in situ hybridization (dFISH) method optimized for detecting two mRNAs of interest in fresh frozen brain sections. Our group has successfully used this approach to study gene co-regulation. More specifically, we have used this dFISH method to explore the anatomical organization, neurochemical properties, and the impact of sensory experience in central sensory circuits, at single cell resolution. This protocol has been validated in brain tissue from mice, rats and songbirds but is expected to be easily adaptable to other vertebrate species, as well as to an array of non-neural tissues. In this film we provide a detailed demonstration of the main steps of this procedure.

Double Fluorescence in situ Hybridization in Fresh Brain Sections

This protocol involves a non-radioactive in-situ hybridization procedure that enables the simultaneous identification of two transcript species, at a single cell resolution, in thin sections of the vertebrate brain.

Here we describe a modified version of a double fluorescence in situ hybridization (dFISH) method optimized for detecting two mRNAs of interest in fresh ...

Methods for Study of Neuronal Morphogenesis: Ex vivo RNAi Electroporation in Embryonic Murine Cerebral Cortex

The cerebral cortex directs higher cognitive functions. This six layered structure is generated in an inside-first, outside-last manner, in which the first born neurons remain closer to the ventricle while the last born neurons migrate past the first born neurons towards the surface of the brain1. In addition to neuronal migration2, a key process for normal cortical function is the regulation of neuronal morphogenesis3. While neuronal morphogenesis can be studied in vitro in primary cultures, there is much to be learned from how these processes are regulated in tissue environments.
We describe techniques to analyze neuronal migration and/or morphogenesis in organotypic slices of the cerebral cortex4,6. A pSilencer modified vector is used which contains both a U6 promoter that drives the double stranded hairpin RNA and a separate expression cassette that encodes GFP protein driven by a CMV promoter7-9. Our approach allows for the rapid assessment of defects in neurite outgrowth upon specific knockdown of candidate genes and has been successfully used in a screen for regulators of neurite outgrowth8. Because only a subset of cells will express the RNAi constructs, the organotypic slices allow for a mosaic analysis of the potential phenotypes. Moreover, because this analysis is done in a near approximation of the in vivo environment, it provides a low cost and rapid alternative to the generation of transgenic or knockout animals for genes of unknown cortical function. Finally, in comparison with in vivo electroporation technology, the success of ex vivo electroporation experiments is not dependant upon proficient surgery skill development and can be performed with a shorter training time and skill.

Methods for Study of Neuronal Morphogenesis: Ex vivo RNAi Electroporation in Embryonic Murine Cerebral Cortex

To conduct a rapid assessment of the function of genes in the development of cerebral cortex, we describe methods involving the ex vivo electroporation of plasmids co-expressing inhibitory RNA (RNAi) and GFP in murine embryonic cortex. This protocol is amenable to the study of various aspects of neurodevelopment such as neurogenesis, neuronal migration and neuronal morphogenesis including dendrite and axon outgrowth.

The cerebral cortex directs higher cognitive functions. This six layered structure is generated in an inside-first, outside-last manner, in which the ...

Fiber-optic Implantation for Chronic Optogenetic Stimulation of Brain Tissue

Elucidating patterns of neuronal connectivity has been a challenge for both clinical and basic neuroscience. Electrophysiology has been the gold standard for analyzing patterns of synaptic connectivity, but paired electrophysiological recordings can be both cumbersome and experimentally limiting. The development of optogenetics has introduced an elegant method to stimulate neurons and circuits, both in vitro1 and in vivo2,3. By exploiting cell-type specific promoter activity to drive opsin expression in discrete neuronal populations, one can precisely stimulate genetically defined neuronal subtypes in distinct circuits4-6. Well described methods to stimulate neurons, including electrical stimulation and/or pharmacological manipulations, are often cell-type indiscriminate, invasive, and can damage surrounding tissues. These limitations could alter normal synaptic function and/or circuit behavior. In addition, due to the nature of the manipulation, the current methods are often acute and terminal. Optogenetics affords the ability to stimulate neurons in a relatively innocuous manner, and in genetically targeted neurons. The majority of studies involving in vivo optogenetics currently use a optical fiber guided through an implanted cannula6,7; however, limitations of this method include damaged brain tissue with repeated insertion of an optical fiber, and potential breakage of the fiber inside the cannula. Given the burgeoning field of optogenetics, a more reliable method of chronic stimulation is necessary to facilitate long-term studies with minimal collateral tissue damage. Here we provide our modified protocol as a video article to complement the method effectively and elegantly described in Sparta et al.8 for the fabrication of a fiber optic implant and its permanent fixation onto the cranium of anesthetized mice, as well as the assembly of the fiber optic coupler connecting the implant to a light source. The implant, connected with optical fibers to a solid-state laser, allows for an efficient method to chronically photostimulate functional neuronal circuitry with less tissue damage9 using small, detachable, tethers. Permanent fixation of the fiber optic implants provides consistent, long-term in vivo optogenetic studies of neuronal circuits in awake, behaving mice10 with minimal tissue damage.

The development of optogenetics now provides the means to precisely stimulate genetically defined neurons and circuits, both in vitro and in vivo. Here we describe the assembly and implantation of a fiber optic for chronic photostimulation of brain tissue.

Elucidating patterns of neuronal connectivity has been a challenge for both clinical and basic neuroscience. Electrophysiology has been the gold standard ...

Isolation of Cerebrospinal Fluid from Rodent Embryos for use with Dissected Cerebral Cortical Explants

The CSF is a complex fluid with a dynamically varying proteome throughout development and in adulthood. During embryonic development, the nascent CSF differentiates from the amniotic fluid upon closure of the anterior neural tube. CSF volume then increases over subsequent days as the neuroepithelial progenitor cells lining the ventricles and the choroid plexus generate CSF. The embryonic CSF contacts the apical, ventricular surface of the neural stem cells of the developing brain and spinal cord. CSF provides crucial fluid pressure for the expansion of the developing brain and distributes important growth promoting factors to neural progenitor cells in a temporally-specific manner. To investigate the function of the CSF, it is important to isolate pure samples of embryonic CSF without contamination from blood or the developing telencephalic tissue. Here, we describe a technique to isolate relatively pure samples of ventricular embryonic CSF that can be used for a wide range of experimental assays including mass spectrometry, protein electrophoresis, and cell and primary explant culture. We demonstrate how to dissect and culture cortical explants on porous polycarbonate membranes in order to grow developing cortical tissue with reduced volumes of media or CSF. With this method, experiments can be performed using CSF from varying ages or conditions to investigate the biological activity of the CSF proteome on target cells.

The ventricular cerebrospinal fluid (CSF) bathes the neuroepithelial and cerebral cortical progenitor cells during early brain development in the embryo. Here we describe the method developed to isolate ventricular CSF from rodent embryos of different ages in order to investigate its biological function. In addition, we demonstrate our cerebral cortical explant dissection and culture technique that allows for explant growth with minimal volumes of culture medium or CSF.

The CSF is a complex fluid with a dynamically varying proteome throughout development and in adulthood. During embryonic development, the nascent CSF ...

Isolation and Culture of Endothelial Cells from the Embryonic Forebrain

Embryonic brain endothelial cells can serve as an important tool in the study of angiogenesis and neurovascular development and interactions. The two vascular networks of the embryonic forebrain, pial and periventricular, are spatially distinctive and have different origins and growth patterns. Endothelial cells from the pial and periventricular vascular networks have unique gene expression profiles and functions. Here we present a step-by-step protocol for isolation, culture, and verification of pure populations of endothelial cells from the periventricular vascular network (PVECs) of the embryonic forebrain (telencephalon). In this approach, telencephalon devoid of pial membrane obtained from embryonic day 15 mice is minced, digested with collagenase/dispase, and dispersed mechanically into a single cell suspension. PVECs are purified from cell suspension using positive selection with anti-CD-31/PECAM-1 antibody conjugated to MicroBeads using a strong magnetic separation method. Purified cells are cultured on collagen 1&#160;coated culture dishes in endothelial cell culture medium until they become confluent and further subcultured. PVECs obtained with this protocol exhibit cobblestone and spindle shaped phenotypes, as visualized by phase-contrast light microscopy and fluorescence microscopy. Purity of PVEC cultures was established with endothelial cell markers. In our hands, this method reliably and consistently yields pure populations of PVECs. This protocol will benefit studies aimed at gaining mechanistic insights into forebrain angiogenesis, understanding PVEC interactions, and cross-talks with neuronal cell types and holds tremendous potential for therapeutic angiogenesis.

This video demonstrates an easy and reliable strategy for preparation of pure cultures of endothelial cells from the embryonic forebrain within 10-12 days and will be useful for research focused on many aspects of cerebral angiogenesis.

Embryonic brain endothelial cells can serve as an important tool in the study of angiogenesis and neurovascular development and interactions. The two ...

Isolation and Cannulation of Cerebral Parenchymal Arterioles

Intracerebral parenchymal arterioles (PAs), which include parenchymal arterioles, penetrating arterioles and pre-capillary arterioles, are high resistance blood vessels branching out from pial arteries and arterioles and diving into the brain parenchyma. Individual PA perfuse a discrete cylindrical territory of the parenchyma and the neurons contained within. These arterioles are a central player in the regulation of cerebral blood flow both globally (cerebrovascular autoregulation) and locally (functional hyperemia). PAs are part of the neurovascular unit, a structure that matches regional blood flow to metabolic activity within the brain and also includes neurons, interneurons, and astrocytes. Perfusion through PAs is directly linked to the activity of neurons in that particular territory and increases in neuronal metabolism lead to an augmentation in local perfusion caused by dilation of the feed PA. Regulation of PAs differs from that of better-characterized pial arteries. Pressure-induced vasoconstriction is greater in PAs and vasodilatory mechanisms vary. In addition, PAs do not receive extrinsic innervation from perivascular nerves &#8212; innervation is intrinsic and indirect in nature through contact with astrocytic endfeet. Thus, data regarding contractile regulation accumulated by studies using pial arteries does not directly translate to understanding PA function. Further, it remains undetermined how pathological states, such as hypertension and diabetes, affect PA structure and reactivity. This knowledge gap is in part a consequence of the technical difficulties pertaining to PA isolation and cannulation. In this manuscript we present a protocol for isolation and cannulation of rodent PAs. Further, we show examples of experiments that can be performed with these arterioles, including agonist-induced constriction and myogenic reactivity. Although the focus of this manuscript is on PA cannulation and pressure myography, isolated PAs can also be used for biochemical, biophysical, molecular, and imaging studies.

This manuscript describes a simple and reproducible protocol for isolation of intracerebral arterioles (a group of blood vessels encompassing parenchymal arterioles, penetrating arterioles and pre-capillary arterioles) from mice, to be used in pressure myography, immunofluorescence, biochemistry, and molecular studies.

Intracerebral parenchymal arterioles (PAs), which include parenchymal arterioles, penetrating arterioles and pre-capillary arterioles, are high resistance ...

Fluorescence Activated Cell Sorting (FACS) and Gene Expression Analysis of Fos-expressing Neurons from Fresh and Frozen Rat Brain Tissue

The study of neuroplasticity and molecular alterations in learned behaviors is switching from the study of whole brain regions to the study of specific sets of sparsely distributed activated neurons called neuronal ensembles that mediate learned associations. Fluorescence Activated Cell Sorting (FACS) has recently been optimized for adult rat brain tissue and allowed isolation of activated neurons using antibodies against the neuronal marker NeuN and Fos protein, a marker of strongly activated neurons. Until now, Fos-expressing neurons and other cell types were isolated from fresh tissue, which entailed long processing days and allowed very limited numbers of brain samples to be assessed after lengthy and complex behavioral procedures. Here we found that yields of Fos-expressing neurons and Fos mRNA from dorsal striatum were similar between freshly dissected tissue and tissue frozen at -80 &#186;C for 3 - 21 days. In addition, we confirmed the phenotype of the NeuN-positive and NeuN-negative sorted cells by assessing gene expression of neuronal (NeuN), astrocytic (GFAP), oligodendrocytic (Oligo2) and microgial (Iba1) markers, which indicates that frozen tissue can also be used for FACS isolation of glial cell types. Overall, it is possible to collect, dissect and freeze brain tissue for multiple FACS sessions. This maximizes the amount of data obtained from valuable animal subjects that have often undergone long and complex behavioral procedures.

Fluorescence Activated Cell Sorting FACS and Gene Expression Analysis of Fos-expressing Neurons from Fresh and Frozen Rat Brain Tissue

Here we present a Fluorescence Activated Cell Sorting (FACS) protocol to study molecular alterations in Fos-expressing neuronal ensembles from both fresh and frozen brain tissue. The use of frozen tissue allows FACS isolation of many brain areas over multiple sessions to maximize the use of valuable animal subjects.

The study of neuroplasticity and molecular alterations in learned behaviors is switching from the study of whole brain regions to the study of specific ...

A Rat Model of Mild Intrauterine Hypoperfusion with Microcoil Stenosis

Intrauterine hypoperfusion/ischemia is one of the major causes of intrauterine/fetal growth restriction, preterm birth, and low birth weight. Most studies of this phenomenon have been performed in either models with severe intrauterine ischemia or models with gradient degree of intrauterine hypoperfusion. No study has been performed in a model on uniform mild intrauterine hypoperfusion (MIUH). Two models have been used for studies of MIUH: a model based on suture ligation of either side of the arterial arcade formed with the uterine and ovarian arteries, and a transient model based on clipping the bilateral ovarian arteries and aorta having patency. Those two rodent models of MIUH have some limitations, e.g., not all fetuses are subjected to MIUH, depending on their position in the uterine horn. In our MIUH model, all fetuses are subjected to a comparable level of intrauterine hypoperfusion. MIUH was achieved by mild stenosis of all four arteries feeding the uterus, i.e., the bilateral uterine and ovarian arteries.
Arterial stenosis was induced by metal microcoils wrapped around the feeding arteries. Producing arterial stenosis with microcoils allowed us to control, optimize, and reproduce decreased blood flow with very little inter-animal variability and a low mortality rate, thus enabling accurate evaluation. When microcoils with an inner diameter of 0.24 mm were used, the blood flow in both the placenta and fetus was mildly decreased (approximately 30% from the pre-stenosis level in the placenta). The offspring of our MIUH model clearly demonstrates long-lasting alterations in neurological, neuroanatomical and behavioral test results.

Mild intrauterine hypoperfusion was produced by artery stenosis with metal microcoils wrapped around the uterine and ovarian arteries in rats at embryonic day 17. This procedure produced prenatal hypoperfusion and intrauterine growth restriction.

Intrauterine hypoperfusion/ischemia is one of the major causes of intrauterine/fetal growth restriction, preterm birth, and low birth weight. Most studies ...

Isolation and RNA Extraction of Neurons, Macrophages and Microglia from Larval Zebrafish Brains

To gain a detailed understanding of the role of different CNS cells during development or the establishment and progression of brain pathologies, it is important to isolate these cells without changing their gene expression profile. The zebrafish model provides a large number of transgenic fish lines in which specific cell types are labelled; for example neurons in the NBT:DsRed line or macrophages/microglia in the mpeg1:eGFP line. Furthermore, antibodies have been developed to stain specific cells, such as microglia with the 4C4 antibody.
Here, we describe the isolation of neurons, macrophages and microglia from larval zebrafish brains. Central to this protocol is the avoidance of an enzymatic tissue digestion at 37 &#176;C, which could modify cellular profiles. Instead a mechanical system of tissue homogenization at 4 &#176;C is used. This protocol entails homogenization of brains into cell suspension, their immuno-staining and the isolation of neurons, macrophages and microglia by FACS. Afterwards, we extracted RNA from those cells and evaluated their quality/quantity. We managed to obtain RNA of high quality (RNA Integrity Number (RIN) &#62; 7) to perform qPCR on macrophages/microglia and neurons, and transcriptomic analysis on microglia. This approach enables a better characterization of these cells, as well as a clearer understanding about their role in development and pathologies.

We present a protocol to isolate neurons, macrophages and microglia from larval zebrafish brains under physiological and pathological conditions. Upon isolation, RNA is extracted from these cells to analyze their gene expression profile. This protocol allows for the collection of high-quality RNA for performing downstream analysis like qPCR and transcriptomics.

To gain a detailed understanding of the role of different CNS cells during development or the establishment and progression of brain pathologies, it is ...

Optimization of Laser-Capture Microdissection for the Isolation of Enteric Ganglia from Fresh-Frozen Human Tissue

The purpose of this method is to obtain high-integrity RNA samples from enteric ganglia collected from unfixed, freshly-resected human intestinal tissue using laser capture microdissection (LCM). We have identified five steps in the workflow that are crucial for obtaining RNA isolates from enteric ganglia with sufficiently high quality and quantity for RNA-seq. First, when preparing intestinal tissue, each sample must have all excess liquid removed by blotting prior to flattening the serosa as much as possible across the bottom of large base molds. Samples are then quickly frozen atop a slurry of dry ice and 2-methylbutane. Second, when sectioning the tissue, it is important to position cryomolds so that intestinal sections parallel the full plane of the myenteric plexus, thereby yielding the greatest surface area of enteric ganglia per slide. Third, during LCM, polyethylene napthalate (PEN)-membrane slides offer the greatest speed and flexibility in outlining the non-uniform shapes of enteric ganglia when collecting enteric ganglia. Fourth, for distinct visualization of enteric ganglia within sections, ethanol-compatible dyes, like Cresyl Violet, offer excellent preservation of RNA integrity relative to aqueous dyes. Finally, for the extraction of RNA from captured ganglia, we observed differences between commercial RNA extraction kits that yielded superior RNA quantity and quality, while eliminating DNA contamination. Optimization of these factors in the current protocol greatly accelerates the workflow and yields enteric ganglia samples with exceptional RNA quality and quantity.

The goal of this protocol is to obtain high-integrity RNA samples from enteric ganglia isolated from unfixed, freshly-resected human intestinal tissue using laser capture microdissection (LCM). This protocol involves preparing flash-frozen samples of human intestinal tissue, cryosectioning, ethanolic staining and dehydration, LCM, and RNA extraction.

The purpose of this method is to obtain high-integrity RNA samples from enteric ganglia collected from unfixed, freshly-resected human intestinal tissue ...