The here introduced protocol allows characterization of the lung homing capacity of primary human lymphocytes under in vivo inflammatory conditions. Pulmonary infiltration of adoptively transferred human immune cells in a mouse model of allergic inflammation can be imaged and quantified by light-sheet fluorescence microscopy of chemically cleared lung tissue.
Overwhelming tissue accumulation of highly activated immune cells represents a hallmark of various chronic inflammatory diseases and emerged as an attractive therapeutic target in the clinical management of affected patients. In order to further optimize strategies aiming at therapeutic regulation of pathologically imbalanced tissue infiltration of pro-inflammatory immune cells, it will be of particular importance to achieve improved insights into disease- and organ-specific homing properties of peripheral lymphocytes. The here described experimental protocol allows to monitor lung accumulation of fluorescently labeled and adoptively transferred human lymphocytes in the context of papain-induced pulmonary inflammation. In contrast to standard in vitro assays frequently used for the analysis of immune cell migration and chemotaxis, the now introduced in vivo setting takes into account lung-specific aspects of tissue organization and the influence of the complex inflammatory scenario taking place in the living murine organism. Moreover, three-dimensional cross-sectional light-sheet fluorescence microscopic imaging does not only provide quantitative data on infiltrating immune cells, but also depicts the pattern of immune cell localization within the inflamed lung. Overall, we are able to introduce an innovative technique of high value for immunological research in the field of chronic inflammatory lung diseases, which can be easily applied by following the provided step-by-step protocol.
Classic inflammatory disorders of the lung, such as allergic asthma and chronic obstructive pulmonary disease (COPD), are well known to be driven by an increased recruitment of activated lymphocytes into the pulmonary tissue1,2. Lymphocyte-released cytokines (e.g., IL-4, IL-5, IL-9, IL-13, IFN-γ and TNF-α) further promote chemotaxis of innate and adaptive immune cells, induce fibrotic airway remodeling or directly damage the lung parenchyma2. So far, the underlying mechanisms responsible for the pathological accumulation of lymphocytes within lung tissue are not yet fully understood. In analogy to tissue-selective T cell imprinting described for gut and skin homing, pulmonary dendritic cells (DCs) are obviously able to prime peripheral T cells for preferential lung infiltration, at least partly via the induction of CCR4 expression on the surface of lymphocytes3. Besides CCR4, airway-infiltrating T cells are also characterized by a particularly increased expression of the chemokine receptors CCR5 and CXCR3 compared to T cells within the peripheral blood1,4,5. Overall, existing data are consistent with the concept that lung homing of T lymphocytes under physiological or inflammatory conditions involves a number of different chemokine receptors and their respective ligands and thus crucially depends on a closely controlled collaboration between innate and adaptive immune cells1. Especially, during the initial phase of pathogen or allergen exposure, cells of the innate immune system respond to TLR stimulation or to IgE-mediated cross-linking by the immediate release of different chemoattractants, like LTB4, CCL1, CCL17, CCL22, CCL20, CXCL10 and PGD21,6,7. As a prime example, the interaction between PGD2 and the chemoattractant receptor CRTh2 is known to be of particular importance for chemotaxis of Th2 cells and thus appeared as promising therapeutic target in the clinical management of asthma. Indeed, patients with moderate asthma showed an improvement of symptoms and a significant increase of the forced expiratory volume in one second (FEV1) after treatment with a selective CRTh2 antagonist compared to the placebo group8,9. In a more progressed state of the inflammatory response, already recruited T cells are able to further amplify pulmonary lymphocyte accumulation via the release of IL-4 and IL-13 as potent stimuli for pulmonary DCs. Subsequently, these myeloid-derived innate cells up-regulate the expression of CCL17 and CCL22 in a STAT6-dependent manner1,10,11. Although the complexity of the described scenario still hinders a complete understanding of T cell lung homing, it offers a plethora of molecular targets for a potentially optimized therapeutic control of inflammatory or allergic pulmonary diseases. Therefore, there is an urgent need of innovative experimental techniques, which are able to further deepen and complement our knowledge in the field of T cell chemotaxis and lung homing.
Due to the fact that lung homing of lymphocytes within the human body is influenced by multiple cellular, humoral and physical parameters1, most of the existing experimental methods are not able to model the whole complexity of this immunological process. Instead, many standard protocols for the analysis of lung homing selectively focus on a specific aspect involved in the cascade of lymphocyte attraction, adhesion, migration and retention. Besides a purely descriptive determination of the mRNA or protein expression pattern of integrins and chemokine receptors on peripheral or lung-infiltrating lymphocytes and the complementary measurement of respective chemokine levels in blood, bronchoalveolar lavage (BAL) or pulmonary tissue12,13,14,15, well-established in vitro cell culture assays allow a functional characterization of lymphocyte adhesion or chemotaxis upon defined experimental conditions16,17,18. In principle, static in vitro adhesion assays monitor the binding capacity of cultured lymphocytes to an endothelial monolayer or to glass slides coated with recombinant endothelial adhesion molecules (e.g., MAdCAM-1, VCAM-1), while standard in vitro chemotaxis assays are usually applied in order to quantify the ability of lymphocytes to migrate along a chemokine gradient in a transwell system19. Both in vitro settings enable a controlled adjustment and modulation of experimental conditions, but on the other hand lack important variables known to critically impact on in vivo chemotaxis and adhesion of lymphocytes. Predominantly, static cell culture assays disregard the influence of shear forces caused by the permanent blood flow19 and potentially neglect the involvement of the surrounding immunological milieu and interacting non-lymphocyte immune cells, both present in a living organism. In order to overcome these limitations, the interpretation of results acquired in static in vitro chemotaxis or adherence assays need further validation in dynamic adhesion experiments under flow conditions20,21 and in in vivo models of inflammatory organ pathology19. Indeed, important conclusions on the regulation of T cell lung homing under inflammatory or allergic conditions could be drawn from animal studies analyzing genetically modified mice in defined models of different pulmonary diseases3,22,23. The quantitative comparison of lung infiltrating lymphocytes between wildtype mice and mice with a deficiency for a specific gene of interest represents a well-established and broadly used tool for defining the impact of particular cellular pathways or receptors on the disease-driven pattern of T cell distribution. However, in contrast to before discussed in vitro cell culture assays, a study design based on classical animal models lacks the ability to analyze and monitor primary human T cells directly derived from the blood or BAL of patients suffering from an inflammatory lung disease. Thus, it still remains challenging to functionally validate whether a diagnostically specified lung disease is able to imprint human lymphocytes for preferential lung tropism and how far clinical parameters might impact on this scenario. Recently, a very elegant in vivo approach was introduced in the context of inflammatory bowel diseases (IBD), which was able to overcome most of these limitations and opened new avenues for advanced translational studies on intestinal lymphocyte homing24. Taking advantage of protocols for solvent-based tissue clearing followed by cross-sectional light-sheet fluorescence microscopy as a powerful imaging tool, it was possible to visualize the infiltration and distribution of adoptively transferred human T cells in the intestine of colitic immunodeficient mice24. In particular, this experimental setting implemented two main innovations: (1) Primary human immune cells can be analyzed under experimentally defined in vivo conditions; (2) a rather large area of the diseased organ (about 1.5 cm x 1.5 cm) can be imaged in high resolution quality, followed by 3D-reconstruction. Moreover, several recent studies successfully established the use of solvent-based tissue clearing and light-sheet fluorescence microscopy as important tools for advanced lung imaging25,26. In order to benefit from this technological progress in the field of pulmonary immunology, we now adopted the system for analysis of lung homing.
The here presented protocol provides a step-by-step introduction how to purify and fluorescently label primary human T cells for transfer into mice with induced pulmonary inflammation and, moreover, describes in detail the subsequent process of light-sheet fluorescence microscopic imaging, including organ preparation and image processing. Overall, we hope to support future translational studies in the field of inflammatory or allergic lung diseases by introducing a sophisticated, but nevertheless feasible, experimental model for monitoring human lymphocyte lung homing upon in vivo conditions.
Experiments involving animals were performed in accordance with protocols approved by the relevant local authorities in Erlangen (Regierung von Unterfranken, Würzburg, Germany). Mice were housed under specific pathogen-free conditions. The collection of human blood was approved by the local ethical committee and the institutional review board of the University of Erlangen-Nuremberg. Each patient gave written informed consent.
1. Induce Allergic Lung Inflammation in Mice
NOTE: As described in earlier studies27, the following experimental procedure allows inducing allergic airway inflammation in mice and, accordingly, triggers the accumulation of innate and adaptive immune cells in the BAL. The described protocol has been established in C57BL/6J mice, but adoption to other standard inbred strains should be possible.
See Figure 1A for a summary of the in vivo experimental procedure.
2. Purify and Fluorescently label Human Peripheral Blood CD4+ T Cells
NOTE: Process cells under sterile conditions.
3. Adoptively Transfer Human CD4+ T Cells into Papain-exposed Recipient Mmice
NOTE: See Figure 1A for a summary of the in vivo performed experimental procedure. Perform cell transfer one day after the last intranasal administration of papain.
4. Prepare Lung tissue for Light-sheet Fluorescence Microscopy
NOTE: The following steps are performed 3 h after transfer of fluorescently labeled human cells (step 3.2). The described experimental procedures including harvesting of lung tissue, fixation and solvent-based tissue clearing were adapted from currently described protocols24,28.
5. Perform Light-sheet Fluorescence Microscopy of Wwhole Murine Lung Lobes
NOTE: Please see the Table of Materials for details on the light-sheet fluorescence microscope and the corresponding software, on which the following steps are based. Comparable systems by other manufacturers, however, can be used as well with developer-specific modifications of the following protocol. Before starting, get familiar with the microscope-specific operating manual and follow the technical instructions by the responsible person on site.
6. Post-image Processing and Quantification of Lung-accumulated Human Cells
NOTE: Please see Table of Materials for details on the post-imaging software for 3D analysis, on which the following steps are based. However, alternative post-imaging softwares can be used as well.
The presented protocol describes an experimental mouse model, which allows monitoring and quantifying the accumulation of adoptively transferred human T lymphocytes in the lung via light-sheet fluorescence microscopy. Figure 1A provides a schematic overview of the in vivo steps of the experimental schedule. In order to guarantee reliable results, it is of substantial importance to ensure a good quality of the isolated and fluorescently labeled human CD4+ T cells, which will afterwards be transferred into mice. As representatively depicted in Figure 1B,C, the above described procedure for microbead-based cell enrichment and subsequent fluorescence-labeling usually results in a flow cytometrically determined CD4+ T cell purity >95% and successful fluorescence-labeling of all CD4+ T cells. The quality and penetration depth of light-sheet fluorescence microscopy critically depends on an appropriate grade of tissue clearing. As demonstrated in Figure 1D, the here applied protocol for ECi-based tissue clearing was able to guarantee a high level of organ transparency, indicating a successful refractive index matching. Finally, a representative overall result of the described experimental protocol in form of fully processed light-sheet fluorescence microscopy images is demonstrated in Figure 2B,C,D and in the Supplementary Video. The autofluorescent signal (displayed in grey) provides a helpful tool for imaging the anatomic structure of the lung. The red signal represents lung accumulated human CD4+ T cells. Quantification of light-sheet fluorescence microscopy imaging allows determining the overall number of lung accumulated human CD4+ T cells per defined area of inflamed lung tissue. A strategy for quantification of human cell infiltration is illustrated in Figure 2E. The (optional) flow cytometric detection of fluorescently labeled cells within the BAL of recipient mice, as depicted in Figure 2F, can be used as a supplemental technique in order to confirm the successful tissue migration of adoptively transferred CD4+ T cells. Moreover, the preference of transferred human T cells for selective accumulation in inflamed lung tissue in the here described experimental setting was further supported by the fact that human CD4+ T cells could not be retrieved in the intestinal mucosa of recipient animals as depicted in Figure 2B (right panel).
Figure 1: Schematic overview of the in vivo experimental workflow. (A) Allergic lung inflammation was induced in C57BL/6J mice by inhalation of papain (50 µg/mouse) on three consecutive days (d0, d1 and d2). The next day (d3), freshly isolated and fluorescently labeled human CD4+ T cells were adoptively transferred into mice via tail vein injection. After another three hours, mice were sacrificed, followed by in situ perfusion and fixation of the lungs. Finally, lungs were explanted and further analyzed ex vivo by light-sheet fluorescence microscopy. Optionally, BAL can be collected before lung explantation to perform extended ex vivo analyses of successfully extravasated and migrated human cells. (B) Representative histogram confirming the purity of the isolated CD4+ T cell fraction as determined by flow cytometry. Cells stained by an anti-human CD4 antibody or isotype control are displayed in black and grey, respectively. (C) Representative histogram confirming the efficacy of the fluorescence-labeling of human CD4+ cells prior to adoptive transfer. (D) Representative images of a perfused murine lung lobe before and after clearing with ECi. FI, fluorescence intensity. Please click here to view a larger version of this figure.
Figure 2: Representative analysis of the pulmonary accumulation of human CD4+ cells in the context of lung inflammation in vivo. Human CD4+ cells were isolated from peripheral blood using density gradient centrifugation followed by a magnetic microbead-based purification step. Human CD4+ cells were labeled using a red light-excitable cell proliferation dye and injected intravenously into papain-exposed C57BL/6J mice. After three hours, mice were sacrificed to collect and further analyze the lung tissue via light-sheet fluorescence microscopy. (A) Representative overview of a 3D reconstruction of murine, papain-exposed lung tissue (grey) three hours after intravenous injection of labeled human CD4+ cells (red) as recorded by light-sheet fluorescence microscopy. (B) Lung tissue of a recipient mouse three hours after cell transfer shown as overview or high magnification segment (left panel). Retrieved human CD4+ cells could be visualized as red signals and were either depicted in overlay with the autofluorescence signal of lung tissue or as single-channel image. In order to confirm the specificity of the detected signal, control lung tissue without intravenous cell transfer served as negative control (middle panel). In contrast to lung tissue, no labeled human CD4+ cells (red) could be detected in the ileum of the same recipient mouse (right panel). (C) Post image processing allows analysis of single slices of lung tissue as well as generation of 3D reconstructions of the whole organ segment that can be either represented as maximal intensity projection (MIP) or in surface mode. Representative images are depicted. (D) Quantification strategy is illustrated exemplarily in the same lung as already depicted in Figure 2A. Defined cubes of the analyzed lung segment were selected and the number of human CD4+ cells (red) was quantified for each cube. Results of a subsequently performed quantification (events/813 µm x 813 µm x 1,000 µm cube) are indicated as mean ± SEM. (E) As an additional option, flow cytometric detection of fluorescently labeled human T cells in the BAL of recipient animals can be performed in order to confirm the successful tissue migration of accumulated human CD4+ T cells. Please click here to view a larger version of this figure.
Supplementary Video: Representative video sequence showing accumulated human CD4+ cells within the murine lung tissue. 3D reconstruction of the murine lung displaying accumulated human CD4+ cells (red) within the context of the pulmonary tissue (grey) three hours after intravenous injection into the tail vein. Images are shown as MIP in the beginning and in the surface mode in the end. Images were acquired via light-sheet fluorescence microscopy and processed by post-imaging software for 3D analysis. Please click here to view this video. (Right-click to download.)
The here described experimental setting provides the opportunity to monitor the lung homing capacity of primary human immune cells under in vivo inflammatory conditions and thereby relevantly complements classically performed in vitro adhesion and chemotaxis assays. To take into account specific anatomic organ characteristics of the lung, important aspects of immune cell homing (including chemotaxis and cell distribution within the target organ) as well as the clinical relevance and transferability of acquired data, we took advantage of three technical key features: (1) the analysis of primary human cells within a living murine organism; (2) the papain-mediated induction of pulmonary inflammation; and (3) light-sheet fluorescence microscopy of cleared lung tissue.
Although functional studies of adoptively transferred human immune cells within a living murine organism might be limited in some aspects due to potentially relevant incompatibilities in receptor-ligand-interactions between mice and men, the general concept of humanized mouse models has been established successfully as an important experimental tool in the field of translational medicine during the last decades24,32,33,34,35. Compared to classical animal models, studies in humanized mice offer the advantage to directly analyze patient-derived immune cells in an in vivo scenario and thereby to identify functional alterations imprinted by the particular disease32,36,37. Regarding the relevance of potential mismatches between defined receptors on the surface of human immune cells and their respective murine ligands or vice versa, former studies already indicated that human CD4+ T cells are able to interact efficiently with the murine adhesion molecules MAdCAM-1 and VCAM-1, which represent crucial ligands for integrin-mediated lymphocyte homing32. Accordingly, migration of intravascularly transferred human immune cells into murine gut tissue could successfully be blocked in vivo by treatment with the clinically used α4β7 integrin antibody vedolizumab32. However, even in case that a specific receptor-ligand-interaction of relevance might show a complete human/murine mismatch, it will presumably be possible to overcome this limitation by genetic engineering and, for instance, by transgenic overexpression of the respective human ligand, growth factor, cytokine or receptor within the murine organism33,35.
As a significant influence of chronic inflammation on the local secretion of chemokines, the endothelial expression of adhesion molecules and the subsequent accumulation of lymphocytes has been described in numerous publications38,39,40, the choice of a suitable experimental model of pulmonary inflammation represented a critical step while establishing the above described protocol and might be adapted dependent on the clinical context of each individual study. The here selected setting of papain-induced allergic lung inflammation represents a well-described experimental model, which is based on the capacity of the locally administered cysteine protease papain to irritate the airway epithelium and trigger the subsequent release of alarmins27,41,42. Interestingly, accidental exposition to papain is known to cause asthma development in humans as well43. Dependent on the inhaled dose of papain, exposed mice show an accumulation of innate and adaptive immune cells and elevated levels of type 2 cytokines in the lung27. While dose escalation turned out to result in an enlargement of airspaces (classic histological phenomenon of COPD) and a destruction of blood vessel walls with subsequent hemorrhage, moderate dosing schedules went along with a prominent pulmonary eosinophilia and thus mimicked an important histological feature of human asthma27. As our purpose was to use this experimental model to generate an inflammatory context for the analysis of pulmonary immune cell homing, we carefully tried to avoid papain-induced damage of blood vessels, which might otherwise result in an unphysiological extravasation of human immune cells due to disturbed endothelial integrity. Accordingly, we selected an intermediate dose of 50 µg of papain per mouse per day in the above described protocol. Although development of papain-induced airway inflammation also occurs in the absence of B and T lymphocytes, lung infiltrating adaptive immune cells are known to impact significantly on the course of disease27,42. For instance, regulatory T cells could be identified as potent regulators of papain-induced airway eosinophilia27. In perspective, the active involvement of pulmonary lymphocytes in the inflammatory pathogenesis of papain-exposed mice implicates that the above described protocol might in addition enable to analyze the functional capacity of adoptively transferred and successfully lung-accumulated human immune cells to modulate lung pathology (e.g., via flow cytometric acquisition of eosinophilic counts in BAL). Furthermore, an additionally performed intranasal administration of selected disease-relevant human chemokines just prior to intravascular cell transfer might allow to analyze the impact of these humoral mediators on the lung homing process of distinct human immune cell populations in an in vivo setting. Likewise, the effect of inhibitors on the lung homing process and, subsequently, on inflammatory lung pathology can be studied.
A particular advantage of the here introduced experimental procedure is the precise tracking and localization of lung infiltrating human immune cells by the high-end imaging quality of light-sheet fluorescence microscopy. Recent studies already demonstrated that the technique of light-sheet fluorescence microscopy represents a valuable experimental tool for analysis of intestinal immune cell homing in translational IBD research and is able to overcome the main limitations of conventional immunofluorescence microscopy and flow cytometry24,32. While analyses based on conventional microscopy are usually restricted to a very small and potentially not representative area of the organ and flow cytometry does not take into account the aspect of tissue organization at all, light-sheet fluorescence microscopy of chemically cleared tissue allows an increased penetration depth without major loss of resolution and thus enables a 3D reconstruction of rather large organ sections (up to 1.5 cm x 1.5 cm)24,28,44. For sure, the quality and validity of acquired 3D reconstructions critically depend on the performance of tissue clearance. In the here described setting, we included a slightly adopted version of the recently published protocol for ECi-based tissue clearing44. Compared to other well established strategies for solvent-based tissue clearing26,45,46, the ECi-based protocol combines the advantages of excellent clearing properties, low toxicity of used reagents and a moderate time requirement44. As the capacity of standard light-sheet fluorescence microscopy to resolve finest tissue structures like alveolar capillaries is still limited even under optimal clearing conditions28, it might somehow be difficult to reliably differentiate between intravascular and already extravasated immune cells. The inclusion of an additional standard fluorescence-based blood vessel staining into the here described protocol would most probably not be able to fully overcome this limitation. Thus, studies with a particular focus on the behavior of immune cells within the pulmonary microcirculation or their diapedesis might preferentially take into account a recently published and very elegant protocol for intravital lung imaging based on 2-photon microscopy47. In this experimental setting, a thoracic window was implanted into anesthetized and ventilated mice, through which the stabilized lung can be monitored by a resonant-scanning 2-photon microscopy module, allowing high-resolution imaging throughout the respiratory cycle upon preserved ventilation and perfusion47,48. This technique has been successfully applied in various studies and was able to visualize for instance intrapulmonary platelet biogenesis and the lung entrance of circulating tumor cells49,50. However, besides the requirement of sophisticated instrumental equipment (e.g., mouse ventilation system) and the need of extensive invasive manipulations in living animals (implantation of the thoracic window and intravital microscopy), the main limitation of this live lung microscopy system is the confined z-axis penetration. The performed lung surface imaging only allows analyzing the outer 100 µm of subpleural lung tissue47,48. Dependent on the scientific context, it might be a valuable option to implement 2-photon microscopy of the explanted lungs as an additional procedure into the above described protocol. Analyzing the same lung first by 2-photon microscopy and afterwards by light-sheet fluorescence microscopy might represent a strategy to obtain a quantitative overview of the distribution and localization of human immune cells within the murine lung (light-sheet fluorescence microscopy) and, at the same time, gain more detailed insights into cellular or microvascular processes (2-photon microscopy). Indeed, 2-photon microscopy of murine tracheal explants represents an established imaging tool for analyzing immune cell behavior in the context of experimentally induced lung inflammation51,52. Instead of visualizing small pulmonary capillaries, the successful extravasation and lung tissue infiltration of the transferred human T cells in our experimental model can also be proven by detecting fluorescently labeled human cells within the BAL of recipient animals via flow cytometry as exemplarily depicted in Figure 2E. In general, analyses of the cellular BAL compartment represent a well-established method to characterize the pulmonary immune cell influx in the context of inflammatory respiratory diseases31. Finally, another flow cytometric strategy for quantifying the extravasation of adoptively transferred immune cells within lung tissue was described by Galkina et al. (2005)53 and could potentially be combined with the here described protocol. In the study by Galkina et al., a single intravenous injection of a fluorescence-conjugated anti-CD8 antibody shortly before resection of the lung resulted in an exclusive and complete labeling of before transferred CD8+ T cells within the lung vascular compartment, while already extravasated CD8+ T cells within the lung interstitium remained unstained. Subsequent analyses of in vivo labeled pulmonary immune cells were performed flow cytometrically after ex vivo digestion of lung tissue53. Of course, the process of tissue digestion means that the anatomic separation between the intra- and extravascular lung compartment is abrogated and, thus, there is a general risk of false-positive labeling of interstitial immune cells due to antibody leakage between both compartments. This risk can only be minimized by performing the tissue digestion in the presence of saturating amounts of unlabeled antibody53 or, potentially, by replacing flow cytometric analysis by light-sheet fluorescence microscopy, which makes it possible to completely avoid the destruction of tissue structure.
In summary, the here introduced combination of in vivo homing of fluorescently labeled primary immune cells and subsequent light-sheet fluorescence microscopy is able to reliably identify, quantify and localize lung accumulated human immune cells in an experimental mouse model of pulmonary inflammation.
The authors have nothing to disclose.
The authors gratefully acknowledge funding by the DFG Collaborative Research Centers SFB 1181 and TRR 241. The Optical Imaging Centre Erlangen (OICE) and in particular Ralf Palmisano, Philipp Tripal and Tina Fraaß (Project Z2 of the DFG CRC 1181) are acknowledged for expert technical support for light-sheet fluorescence microscopic imaging.
Agarose NEEO Ultra | Carl Roth GmbH + Co. KG, Karlsruhe, Germany | 2267.4 | |
AlexaFlour594 anti-human CD45 antibody | BioLegend, San Diego, USA | 304060 | |
Ammonium chloride | Carl Roth GmbH + Co. KG, Karlsruhe, Germany | K2981 | |
Cannula 21 G | Becton, Dickinson and Company, Franklin Lakes, USA | 301300 | |
Cell proliferation dye eflour670 | eBioscience Inc., San Diego, USA | 65-0840-85 | |
CD4 MicroBeads, human | Miltenyi Biotech GmbH, Bergisch-Gladbach, Germany | 130-045-101 | |
EDTA (ethylenediaminetetraacetic acid) | Carl Roth GmbH + Co. KG, Karlsruhe, Germany | 8043.1 | |
Potassium-EDTA blood collection tube, 9 ml | Sarstedt AG & Co., Nümbrecht, Germany | 21066001 | |
Ethly cinnamate (ECi) | Sigma-Aldrich, Steinheim, Germany | 112372-100G | |
Ethanol ≥ 99.5 % (EtOH) | Carl Roth GmbH + Co. KG, Karlsruhe, Germany | 5054.3 | |
FBS (fetal bovine serum) Good Forte | PAN-Biotech GmbH, Aidenbach, Germany | P40-47500 | |
Filter 100 µm | VWR International Germany GmbH, Darmstadt, Germany | 732-2758 | |
Imaris Image Analysis Software 9.0.2 | Bitplane AG, Zurich, Switzerland | n.a. | |
ImspectorPro software | Abberior Instruments GmbH, Göttingen, Germany | n.a. | |
Ketamin | Inresa Arzneimittel GmbH, Freiburg, Germany | 3617KET-V | |
LaVision UltraMicroscope II | LaVision BioTec GmbH, Bielefeld, Germany | n.a. | |
MACS MultiStand | Miltenyi Biotech GmbH, Bergisch-Gladbach, Germany | 130-042-303 | |
Multifly cannula 20 G | Sarstedt AG & Co., Nümbrecht, Germany | 851638035 | |
30 G needle | B. Braun Melsungen AG, Melsungen, Hessen, Germany | 9161502 | |
Neubauer counting chamber | neoLab Migge GmbH, Heidelberg, Germany | C-1003 | |
Pattex Glue | Henkel AG & Co, Düsseldorf, Germany | PSK1C | |
LS column | Miltenyi Biotech GmbH, Bergisch-Gladbach, Germany | 130-042-401 | |
Lymphocyte Separation Media (Density 1,077 g/ml) | anprotec | AC-AF-0018 | |
RPMI medium | (Gibco) Life Technologies GmbH, Darmstadt, Germany | 61870-010 | |
Papain | Merck | 1,071,440,025 | |
PBS Dulbecco (phosphate buffered saline) | Biochrom GmbH, Berlin, Germany | L182-10 | |
PerCP/Cy5.5 anti-human CD4 | BioLegend, San Diego, USA | 317428 | |
PerCP/Cy5.5 mouse IgG2b, κ isotype Ctrl | BioLegend, San Diego, USA | 400337 | |
PFA (paraformaldehyde) | Carl Roth GmbH + Co. KG, Karlsruhe, Germany | 0335.1 | |
Potassium hydrogen carbonate | Carl Roth GmbH + Co. KG, Karlsruhe, Germany | P7481 | |
Serological pipette 10 ml | Sarstedt AG & Co., Nümbrecht, Germany | 86.1254.001 | |
Syringe 1 ml | B. Braun Melsungen AG, Melsungen, Hessen, Germany | 9166017V | |
Syringe 5 ml | Becton, Dickinson and Company, Franklin Lakes, USA | 260067 | |
Syringe 20 ml | Becton, Dickinson and Company, Franklin Lakes, USA | 260069 | |
Tube 1.5 ml | Sarstedt AG & Co., Nümbrecht, Germany | 72,706,400 | |
Tube 2 ml | Sarstedt AG & Co., Nümbrecht, Germany | 72.695.400 | |
Tube 2 ml, brown | Sarstedt AG & Co., Nümbrecht, Germany | 72,695,001 | |
Tube 15 ml | Sarstedt AG & Co., Nümbrecht, Germany | 62.554.502 | |
Tube 50 ml | Sarstedt AG & Co., Nümbrecht, Germany | 62.547.254 | |
QuadroMACS Separator | Miltenyi Biotech GmbH, Bergisch-Gladbach, Germany | 130-090-976 | |
Xylazin (Rompun 2%) | Bayer Vital GmbH, Leverkusen, Germany | KPOBD32 |