Animal procedures were performed at the University of Minnesota in facilities accredited by the Association for Assessment and Accreditation of Laboratory Animal Care (AAALAC) and in accordance with protocols approved by the University of Minnesota IACUC, as well as the principles outlined in the National Institute of Health Guide for the Care and Use of Laboratory animals.
2.2. Preparation of Coverslips
German glass 12mm coverslips (Bellco; 1943-10012) were acid washed in a 1M HCl solution overnight at 55 °C, washed twice for 30 minutes with distilled water and then rinsed 30 minutes each in 50%, 75%, and 90% solutions of ethanol. Following washing, coverslips were dried and maintained in at 225 °C oven until immediately prior to coating. Observations regarding the importance of the source, washing and storage of coverslips for cell adherence, survival, and development are detailed in supplemental table 1
One to two days before cultures were prepared, acid-washed coverslips were transferred to 35mm petri dishes (5 coverslips per 35mm dish) and coated with a mixture of 100μg/ml Poly-D-Lysine (PDL, Sigma; p7886) and 4μg/ml laminin (Invitrogen; 23017015). PDL was prepared by dissolving 100mg of PDL in 0.1M borate buffer (pH 8.5). PDL/borate solution was then sterilized through a 0.2μm filter, aliquoted and stored at −80°. Laminin was thawed on ice, aliquoted and stored at −80°. Immediately prior to coverslip coating, PDL was thawed at 32° and Laminin was thawed on ice. When working with laminin, care must be taken to avoid warming the solution too quickly, which can cause laminin to aggregate, reducing its ability to serve as a substrate for cells. After overlaying 50μl of PDL/laminin on each coverslip, the dishes were sealed with parafilm to prevent evaporation and coverslips were incubated overnight at room temperature. Coverslips were then rinsed three times with sterile water. Each 35mm dish was filled with 2ml of neuronal plating media (see section 2.4) and stored in the tissue culture incubator until needed.
2.3. Dissection of striatal and cortical tissues
presents a flow chart of the culturing protocol. Embryonic day 16 (E16) pregnant mice were euthanized by CO2
asphyxiation and the uterus was removed. Within a sterile dissecting hood, embryos were removed from the uterus and decapitated. Heads were stored in room temperature Ca2+
-free Hank’s Balance Salt Solution with 10mM HEPES (CMF-HBSS, the same solution in which all dissections took place). All dissecting equipment was sterilized in 70% ethanol and was routinely dipped in 70% ethanol during the dissection, especially during transitions from exterior structures to interior structures (i.e. between removing brain from skull and working with isolated brain). These steps were undertaken to maintain the sterile environment and avoid contamination. Routine use of this procedure eliminates the need for antibiotic administration at any point during the preparation or maintenance of the cultures. Brains were removed from the skull using forceps and transferred to a wax bottom 35mm petri dish (5ml of paraffin wax in the bottom of dissecting dishes helps protect the tips of delicate tools). The hemispheres were removed from the cerebral peduncles with a sharp forceps. The meninges were removed and hemispheres were placed medial surface up. A region of cortex, roughly corresponding to the pre-frontal cortex, was cut from the medial side of the anterior aspect of the cortex. The remaining cortex overlying the lateral ventricle was folded away to expose the ventricle and ganglionic eminences. The ganglionic eminences (medial and lateral; lateral being the presumptive striatum (Wichterle et al., 2001
)) were removed using angled forceps to “scoop out” the entire eminence from the lateral ventricle and separate it from the overlying cortex. The cortex and ganglionic eminence regions were transferred to separate CMF-HBSS containing petri dishes. After dissection of all pups, tissue was minced using a clean, sterile razor blade. Videos of the dissection procedure are available upon request.
Schematic illustrating the major steps in the cortical-striatal co-culture procedure.
2.4. Tissue dissociation
Minced cortical and striatal tissue was transferred to separate sterile 15ml conical tubes and allowed to settle to the bottom of the tube. CMF-HBSS was removed and replaced by either papain or trypsin digestion solutions. For papain treatment, minced tissue pieces were resuspended in 5 ml of 20–25 units/ml papain (Worthington, LS003126) solution including 1.1mM EDTA and 5.5mM cysteine in CMF-HBSS and incubated for 30 minutes at 37 °C. Cells were allowed to settle to the bottom of the tube, digestion solution was removed and 5ml of inhibition solution (750μg/ml DNAse1, 10mg/ml BSA, and 10mg/ml Type II-O Trypsin inhibitor) was then added and the incubation continued for 15 minutes. Tissue was then centrifuged at 1000g (~3000rpm) for five minutes, resuspended in neuronal plating media (10mM HEPES, 10mM sodium pyruvate, 0.5mM glutamine, 12.5μM glutamate, 10% Newborn Calf Serum, 0.6% Glucose in EMEM (Minimal Essential Media, plus Earl’s salt)) and triturated no more than ten times with a fire polished Pasteur pipette. For trypsin digestion, 10X Trypsin-EDTA (Sigma-Aldrich, T4174) was added to the cell suspension at a final concentration of 0.25%Trypsin and incubated for 30 minutes at 37 °C with occasional gentle agitation. Tissue was monitored during this period and 3mg/ml DNAse1 (Sigma-Aldrich, DN25) added if tissue pieces appear to be connected by a stringy mass of DNA. After trypsinization, cells were centrifuged at 1000g for five minutes, solution was removed and replaced with neuronal plating media and triturated no more than ten times using sterile flame-polished glass pipettes. For either protease treatment, properly digested tissue should be fully dissociated after 10 triturations. Excessive trituration will reduce cell viability, as will introduction of air/bubbles during the trituration. Following dissociation, cells were counted using trypan blue (to determine cell viability) and a hemocytometer. Cell viability should be ≥ 90%.
2.5. Plating and maintenance of neuronal cultures
Cells were plated at a total density of 2×105 cells/35mm dish. For co-cultures, a ratio of 2 parts striatal to 3 parts cortical cells was used. One to three hours after plating the neuronal plating media was replaced with neuronal growth media (50× B27, 0.5mM glutamine, Neurobasal) that had been conditioned 24–48hrs on confluent glia cultures (see 2.7). Every seven days one half of the media was replaced with new glial conditioned neuronal growth media.
2.6 Plasmid DNAs
Three different types of plasmids were used to test DNA transduction methods: (1) pEGFP-C1 (Clontech) expresses enhanced green fluorescent protein (EGFP) under the control of the CMV promoter, (2) pCAG-EGFP (constructed in the Lanier lab) expresses EGFP under the control of the CMV immediate early enhancer and the chicken beta actin promoter (pCAG, Alexopoulou et al, 2008
), (3) EGFP was cloned into pTRE-tight (Clontech) and co-transfected with pCAG-rtTA-advanced (produced by subcloning the rtTA-advanced tet transactivator from pTet-On-Advanced (Clontech) into pCAG). Additional details about plasmid construction are available on request.
For some experiments, striatal cells were electroporated using the Lonza Nucleofector system and the mouse neuron transfection reagent (cat. VAPG-1001). We have found that best results are obtained if the two parts of the transfection solution are mixed on the day of transfection (rather than mixed in advance as per the manufacturer’s instructions) and if the transfection kit is used within 3 months of purchase. For this reason, we buy the smaller kit (10 reactions) unless we plan to do more than 10 transfections in 2 months time. For transfection, 1×106 dissociated striatal cells were transferred to a microfuge tube and centrifuged at 1000g for 5 minutes, then plating media was removed and cells were gently resuspended in 100μl of complete transfection reagent containing 10–12μg of pCAG-EGFP or pEGFP-C1 or a mix of 10μg of pTRE-tight-EGFP and 17μg of pCAG-rtTA plasmid DNA per 1 million cells (keep total DNA volume < 10 μl). Following electroporation, pre-warmed plating media was added to the cuvette, cells were quickly transferred to 2ml of pre-warmed plating media and samples were taken for hemocytometer counting using trypan blue. Cell viability should be ≥90%. We have found that the inclusion of unelectroporated neurons at plating helps the survival and development of electroporated neurons. Therefore, in the co-culture condition, cortical cells were unelectroporated and the striatal cells were electroporated. In the mono-culture condition electroporated striatal cells are mixed 1:1 with unelectroporated striatal neurons at plating.
2.8. Glia Cultures
Glia cultures were prepared from cortices of postnatal (P1-2) mice with the striatal tissue removed. Minced tissue was incubated in CMF-HBSS plus 0.25% trypsin-EDTA and DNAse (either 3mg/ml DNAseI or 1μl/ml Benzonase (Novagen, 70664-3)) at 37°C for 30 minutes. After trypsinization, an equal volume of glia plating media (EMEM, 10mM HEPES, 1mM Sodium Pyruvate, 0.2mM glutamine, 10% NCS, 0.6% Glucose, 1× Penicillin-Streptomycin) was added to inhibit the trypsin and the tissue was collected by centrifugation (1000g for 2 minutes). Tissue was resuspended in glia plating media, mechanically dissociated using a flame-polished glass pipette and filtered through a 0.7μm cell strainer (BD Biosciences, 352350). Cells from one litter of pups were plated one pup per dish onto 10 cm tissue culture dishes (untreated, because coating with PDL does not increase the yield). Glia plating media was replaced one day after culturing and once per week each subsequent week. The first day after plating dishes tend to have a large amount of debris and significant cell division does not occur until approximately 3 days in vitro (div). After 7–10div plates have reached 70–100% confluence and are suitable for use in the preparation of conditioned media. These cultures will be >90% astroglia and appear as large, flat cells that can be somewhat difficult to view under phase contrast.
Alternatively, confluent dishes of glial cells can be frozen and stored in liquid nitrogen following standard protocols. Briefly, glia cultures are trypsinized, resuspended in an equal volume of glia plating media, centrifuged and resuspended in cold freezing media (MEM/20% NCS/10% DMSO). Glia should be aliquoted one 10 cm dish of cultured cells per freezing vial. Vials are transferred to a styrofoam container to slow freeze overnight at −80, and then transferred to liquid nitrogen. To thaw glia, agitate one vial of cells in a 37° water bath until the contents are just thawed (do not let the vial warm up). Immediately transfer the cell suspension to a tube containing 5 volumes of warm glia plating media, centrifuge and resuspend cells in warm glia plating media. Plate one vial of cells per 10 cm dish. If freezing and storage was done properly, >90% of the cells should survive. It should take about 2 weeks for the glia to reach confluence and be good for conditioning.
Glial-conditioned growth media (GCM) was prepared by removing the glia plating media and replacing with 7–10ml of neuronal growth media for 24–48 hrs. Once glial conditioning of neuronal growth media was complete, the GCM was removed and replaced with glia plating media. GCM is used within 24hrs after it is reclaimed from glial plates and stored in the 37°C/5% CO2 incubator when not in use. Glial plates are typically used only once per week for conditioning media and, if used to condition multiple times in one week, given at least 48hrs in glia plating media between conditioning sessions. Confluent glia plates can be maintained in this fashion for three or more months and are discarded if significant microglial proliferation is observed (microglia appear as bright, refractile cells, usually growing on top of the astroglia).
At 14 and 21div cultures were processed for immunofluorescent imaging. Coverslips were fixed in 4% Paraformaldehyde/PHEM (60mM PIPES pH 7.0, 25mM K-HEPES pH 7.0, 10mM EGTA, 2mM MgCl2)/0.12M sucrose buffered fixative for 15–20 minutes at 4°C. Following fixation, cells were rinsed with Phosphate Buffered Saline (PBS) and blocked in 3% fatty acid free bovine serum albumin (BSA; Roche 03117057001) in PBS for 30 minutes at room temperature or overnight at 4°C. The use of a BSA blocking step before and after permeabilization enhances preservation of fine cell structures, especially cytoskeletal components. In addition, use of BSA, rather than serum, makes the blocking compatible with phospho-epitope specific antibody staining. Cells were permeabilized for 10 minutes at room temperature in 0.2% Triton X-100 in PBS after which coverslips were rinsed for 5 minutes in room temperature PBS. After permeabilization, cells were again blocked in 3% BSA in PBS for a minimum of 15 minutes at room temperature. Coverslips were incubated overnight at 4°C with 50μl of primary antibody mixture per slip in 1% BSA in PBS. The following primary antibodies were used: Rabbit anti-DARPP-32 1:250 (Cell Signaling, 2302), mouse anti-βIII Tubulin 1:2000 (Promega, G7121), mouse anti-SV2 1:100 (DSHB, SV2), and monoclonal mouse anti-PSD-95 1:100 (Chemicon, MAB1598). The protocol described above was used for most combinations of antibodies. The only exception was in staining using the PSD-95 antibody. For best exposure of this epitope, cultures were fixed as described above and then treated for 5 minutes with −20°C Methanol. Following overnight incubation at 4°C with primary antibody, coverslips were washed in room temperature PBS and incubated for 1 hour at room temperature with secondary antibody and phalloidin combinations in 1% BSA in PBS. The following secondary antibodies were used: donkey anti-rabbit and anti-mouse conjugated to AMCA, Texas Red, FITC, TRITC, or Cy5 (Jackson Immunoresearch), all used at 1:100 dilutions. Alexa-488 or -594 phalloidins (Molecular Probes) were diluted 1:200. Following the secondary incubation period, coverslips were washed for 5 minutes in room temperature PBS. For nuclear staining, coverslips were submerged in 1μg/ml DAPI in PBS for 30 seconds and rinsed in room temperature PBS for 5 minutes before mounting. Coverslips were mounted on glass slides with 2.5% 1,4-Diazabicyclo-[2.2.2]Octane, 150mM Tris pH 8.0, and 80% glycerol mountant to reduce photobleaching.
2.10. Imaging, Quantification, and Statistical Analysis
Images for Sholl analysis and antibody localization were collected using a Zeiss Axiovert 200M microscope and Openlab software (Improvision/Perkin Elmer). Image adjustments were made using Photoshop CS3 (Adobe) with brightness and intensity changes standardized for each experiment (all conditions). All image analysis was conducted using one or more plugin for ImageJ (NIH, http://rsbweb.nih.gov/ij/
; detailed below). All statistical analysis was conducted using GraphPad Prism v4.0 (GraphPad Software).
For experiments calculating the percentage of DARPP-32 positive neurons, a minimum of 20 fields per coverslip was imaged at 10× magnification. Exposure times for each channel were standard across conditions. Multi-channel images of the same field were merged into a single “stacked” image in ImageJ and the cell-counter plug-in (http://rsbweb.nih.gov/ij/plugins/cell-counter.html
) was used to count and label DAPI stained nuclei, βIII-Tubulin-positive neurons, and DARPP-32 staining. To be counted as DARPP-32 positive, complete soma staining and a visible staining in the dendrites was required. This inclusion criteria is relatively strict when compared to other reports that require detection of DARPP-32 only in the soma (e.g. Ivkovic and Ehrlich, 1999
). The percentage of DARPP-32 positive cells per field was calculated and each field was treated as an independent observation. Replications from 2 independent cultures were pooled. Percentages were compared between time points using Mann-Whitney U to compare two groups of non-Gaussian distribution, p<0.05 was considered significant.
For experiments investigating the morphological complexity of DARPP-32 positive neurons, Sholl analysis was performed (Sholl, 1953
). All isolated DARPP-32 positive neurons on the coverslip were imaged at 20× magnification using standard exposure time across conditions. The ImageJ concentric rings plugin (http://rsbweb.nih.gov/ij/plugins/concentric-circles.html
) was used to place concentric rings every 10μm out to 150μm from the cell’s center. The cell-counter plugin was used to count and label processes (axons and dendrites) crossing each ring starting 20μm from the center of the cell. Replications from 3 independent cultures were pooled. Crossings were compared across the distance measured and between conditions using a two-way ANOVA with Bonferonni’s post-test to determine distances of significant difference, p<0.05 was considered significant.
For experiments determining spine density and morphology in striatal monoculture and striatal-cortical co-culture, EGFP positive, DARPP-32 positive neurons were imaged at 100× magnification using a standard exposure time across conditions. Images were collected using a Personal Deltavision Microscope and softWoRx software (Applied Precision). Terminal tips of isolated dendrites were identified and z-stack images were collected at 0.15–0.2μm intervals through the dendrite and the stacks were deconvolved using softWoRx software (Applied Precision). The deconvolved tiff files were imported into Neuronstudio (Mount Sinai School of Medicine, Rodriguez et al 2008
) and subjected to semi-automated spine analysis. Densities calculated from individual segments of dendrite were treated as an independent observation and densities calculated from 3 independent culture preps were pooled. For dendritic spine morphology comparisons, a trained analyst, blind to condition, confirmed that spines identified by Neuronstudio met criteria for inclusion (observable neck connected to the dendrite shaft). Neuronstudio collects three parameters used to automatically qualify dendritic spines into morphological categories; spine head diameter (hd), spine neck length (nl), and spine neck diameter (nd). In step one, spines with a neck ratio (hd:nd) greater than 1.1 will be thin or mushroom and spines with a neck ratio less than 1.1 will be thin or stubby. For spines that have been identified as either thin or stubby, spines with a thin ratio (nl:hd) greater than 2.5 will be classified as thin and spines with a thin ratio of less than 2.5 will be classified as stubby. For spines that have been identified as either thin or mushroom, spines with head diameters greater than 0.35μm will be classified as mushroom, spines with head diameters less than 0.35μm are classified as stubby. Occasionally Neuronstudio identified spines but was unable to collect the spine neck diameter. In these cases spines were manually categorized based on the nl:hd ratio and head diameter. Spines were categorized as mushroom if they had a neck length to head diameter ratio of less than 1 and an absolute head diameter greater than 0.35μm (nl:hd <1, hd >0.35μm). Spines with a neck length to head diameter ratio of greater than 2.5 were classified as thin (nl:hd >2.5) and all other spines were classified as stubby (nl:hd >1, <2.5). Densities and morphological measurements were compared between conditions using Mann-Whitney U to compare two groups of non-Gaussian distribution, p<0.05 was considered significant. Measurements are reported in the results section and displayed in figures as mean ± SEM.
Coverslips were transferred to a recording chamber superfused with artificial cerebral spinofluid (ACSF) at 22–23°C saturated with 95% O2/5% CO2 and containing 119mM NaCl, 2.5mM KCl, 1.0mM NaH2PO4, 1.3mM MgSO4, 2.5mM CaCl2, 26.2mM NaHCO3, and 11mM glucose. Picrotoxin (100 mM) was added to block GABA-A receptor-mediated IPSCs (inhibitory post-synaptic currents). Cells were visualized using infrared-differential interference contrast (DIC) optics. MSNs were identified by their morphology (i.e. soma size, dendrite organization, dendritic spines) and hyperpolarized resting membrane potential. In some cultures, DIC observations were confirmed by EGFP expression (, left panel)
Figure 5 Basic electrophysiological characterization of mono-and co-cultured MSNs. (A) Analysis of synaptic transmission. Left, co-cultured MSNs (CO, n = 7 cells) exhibit a significantly greater mEPSC frequency and lower amplitude compared to mono-cultured MSNs (more ...)
To assess excitatory synaptic transmission, neurons were voltage clamped at −70 mV using a Multiclamp 700A amplifier (Molecular Devices, Foster City, CA) and miniature EPSCs (excitatory post-synaptic currents) were collected in the presence of lidocaine hydrochloride (1mM). Electrodes (3–5 MΩ) contained 117mM cesium gluconate, 2.8mM NaCl, 20mM HEPES, 0.4mM EGTA, 5mM tetraethylammonium-Cl, 2mM MgATP, and 0.3mM MgGTP, pH 7.2–7.4 (285–295 mOsm). Series resistance (10–30 MΩ) and input resistance were monitored online with a 4 mV depolarizing step (100 ms) at 0.1 Hz. Data were low-pass filtered at 2 kHz, digitized at 10 kHz, and collected and analyzed using custom software (Igor Pro; Wavemetrics, Lake Oswego, OR). Quantal events were analyzed using Minianalysis software (Synaptosoft, Decatur, GA) and verified by eye.
To assess firing properties, kynurenic acid (2mM) was used to block glutamatergic transmission during recording. Whole-cell current-clamp recordings were performed with electrodes (3–5 MΩ) containing 120mM K-gluconate, 20mM KCl, 10mM HEPES, 0.2mM EGTA, 2mM MgCl2, 4mM Na2ATP, and 0.3mM Tris–GTP. Data were low-pass filtered at 5 kHz, digitized at 10 kHz, and collected and analyzed using custom software (Igor Pro; Wavemetrics). Membrane potentials were held at approximately −70 mV. Series resistances ranged from 10 to 18MΩ and input resistances (Ri) were monitored on-line with a +40pA, 150ms current injection given before every 800ms current injection stimulus. Firing was obtained through a series of hyperpolarizing and depolarizing current injections (800 ms duration at 0.1 Hz, −20 to +260pA range with a 20pA step increment). Resting membrane potentials were corrected for liquid junction potential (~ 14mV).
The mEPSC amplitudes and frequencies are presented as mean ± SEM in the results section and shown in a box-and-whisker plot in . The box-and-whisker plot shows maximum, upper quartile, median, lower quartile and minimum. Statistical analyses were performed using GraphPad Prism 5 (GraphPad Software; La Jolla, CA). Statistical significance was assessed using two-tailed Student’s t tests, p<0.05 was considered significant.