Neurospheres are primary cell aggregates that comprise neural stem cells and progenitor cells. These 3D structures are an excellent tool to determine the differentiation and proliferation potential of neural stem cells, as well as to generate cell lines than can be assayed over time. Also, neurospheres can create a niche (in vitro) that allows the modeling of the dynamic changing environment, such as varying growth factors, hormones, neurotransmitters, among others. Microtus ochrogaster (prairie vole) is a unique model for understanding the neurobiological basis of socio-sexual behaviors and social cognition. However, the cellular mechanisms involved in these behaviors are not well known. The protocol aims to obtain neural progenitor cells from the neurogenic niches of the adult prairie vole, which are cultured under non-adherent conditions, to generate neurospheres. The size and number of neurospheres depend on the region (subventricular zone or dentate gyrus) and sex of the prairie vole. This method is a remarkable tool to study sex-dependent differences in neurogenic niches in vitro and the neuroplasticity changes associated with social behaviors such as pair bonding and biparental care. Also, cognitive conditions that entail deficits in social interactions (autism spectrum disorders and schizophrenia) could be examined.
The prairie vole (Microtus ochrogaster), a member of the Cricetidae family, is a small mammal whose life strategy develops as a socially monogamous and highly sociable species. Both males and females establish an enduring pair bond after mating or long periods of cohabitation characterized by sharing the nest, defending their territory, and displaying biparental care for their progeny1,2,3,4. Thus, the prairie vole is a valuable model for understanding the neurobiological basis of socio-sexual behavior and impairments in social cognition5.
Adult neurogenesis is one of the most paramount processes of neural plasticity that leads to behavioral changes. For example, our research group reported in male voles that social cohabitation with mating increased cell proliferation in the subventricular zone (VZ) and subgranular zone in the dentate gyrus (DG) of the hippocampus, suggesting that adult neurogenesis can play a role in the formation of pair bonding induced by mating in prairie voles (unpublished data). On the other hand, although the brain regions where new neurons are generated and integrated are well known, the molecular and cellular mechanisms involved in these processes remain undetermined due to technical drawbacks in the whole brain model6. For instance, the signaling pathways controlling gene expression and other cellular activities have a relatively short activation period (detection of phosphoproteome)7. One alternative model is isolated and cultured adult neural stem cells or progenitor cells to elucidate molecular components involved in adult neurogenesis.
The first approach to maintain in vitro neural precursors from adult mammal (mouse) brain was the assay of neurospheres, which are cellular aggregates growing under non-adherent conditions which preserve their multipotent potential to generate neurons, as well as astrocytes8,9,10. During their development, there is a selection process where only the precursors will respond to mitogens such as the Epidermal Growth Factor (EGF) and Fibroblast Growth Factor 2 (FGF2) to proliferate and generate neurospheres8,9,10.
To our knowledge, no protocol is reported in the literature to obtain adult neural progenitors from prairie voles. Here, we established the culture conditions to isolate neuronal progenitors from neurogenic niches and their in vitro maintenance through the neurosphere formation assay. Thus, experiments can be designed to identify the molecular and cellular mechanisms involved in proliferation, migration, differentiation and survival of the neural stem cells and progenitors, processes that are still unknown in the prairie vole. Moreover, elucidating in vitro differences in the properties of the cells derived from the VZ and DG could provide information about the role of neurogenic niches in neural plasticity associated with changes in socio-sexual behavior and cognitive behaviors, and deficits in social interactions (autism spectrum disorder and schizophrenia), which could also be sex-dependent.
The study was approved by the Research Ethics Committee of the Instituto de Neurobiología, Universidad Nacional Autónoma de México, Mexico and Instituto Nacional de Perinatologia (2018-1-163). The reproduction, care and humane endpoints of the animals were established following the Official Mexican Standard (NOM-062-Z00-1999) based on the “Ley General de Salud en Materia de Investigación para la Salud” (General Health Law for Health Research) of the Mexican Secretaria of Health.
1. Solutions and stocks preparation
- Prepare an N2 culture medium with 485 mL of Dulbecco's Modified Eagle Medium-F12 (DMEM-F12), 5 mL of N2 supplement (100x), 5 mL of glutamine supplement (100x) and 5 mL of antibiotic-antimycotic (100x).
- Prepare a B27 culture medium with 480 mL of Neurobasal medium, 10 mL of B27 supplement (50x), 5 mL of glutamine supplement, and 5 mL of antibiotic-antimycotic (100x).
- Reconstitute collagenase powder in 1x PBS (Phosphate-buffered saline) to obtain aliquots with an activity of 100 units/µL (1000x) and store at -20 °C. Notice, collagenase activity depends on the lot number of the companies.
- Prepare dispase stock aliquots by dissolving 5 mg of dispase powder in 1x PBS (50 mg/mL). Store at -20 °C.
- Prepare an enzymatic solution with 100 mL of DMEM-F12 medium, 50 µL of stock collagenase (100 units/µL) to have a final concentration of 50 U/mL and 333 µL of stock dispase (50 mg/mL) to have a final concentration of 0.33 mg/mL.
- To prepare a washing solution, to 1,000 mL of 1x PBS, add 0.4766 g of HEPES (final concentration 2 mM), 3.6 g of D-glucose (final concentration 20 mM) and 2.1 g of NaHCO3 (final concentration 25 mM).
- Prepare poly-L-ornithine stock aliquots (1 mg/mL) using sterile water and store at -20 °C.
- Prepare a working solution of poly-L-ornithine. Dilute a stock aliquot (1mg/mL) in 49 mL of sterile water for a final concentration of 20 µg/mL.
- Prepare a working solution of laminin. Dilute 25 µL of laminin (1 mg/mL original stock) in 5 mL of sterile water for a final concentration of 5 µg/mL.
NOTE: After preparation, filter the culture media, working and stock solutions to avoid contamination. Use a syringe or bottle-top vacuum filters (polyethersulfone membrane with a 0.2 µm pore size). The culture media and work solutions can be stored for up to 30 days at 4 °C, while the stocks can be stored for up to four months at -20 °C.
2. Preparation before starting the microdissection
- Sterilize surgical instruments by autoclaving or with a hot glass bead dry sterilizer.
- Clean the microdissection surface area under strict aseptic and antiseptic conditions (e.g., with ozonized water).
NOTE: The timing of microdissection of both neurogenic niches from each vole brain is approximately 30 min. Working with 1-4 animals for the entire procedure is recommended.
3. Extraction of the whole brain
- Anesthetize the adult vole (12-16 weeks) with an overdose of pentobarbital (6.3 mg/animal) through intraperitoneal injection. Verify the depth of anesthesia by the absence of pedal reflex in response to a firm toe pinch.
- Once the vole is entirely anesthetized, induce euthanasia by decapitation and recover the head.
- Dissect the skin from the skull with scissors, making a caudal-rostral incision (15 mm long) to expose the skull.
- Cut the occipital and interparietal bones and trace an incision into the skull along the sagittal and parietal sutures.
- Make a hole in the skull at the junction of frontal and parietal bones using scissors, being very careful not to damage the brain tissue.
- To expose the brain, remove the remaining cranium fragments that cover both brain hemispheres with sharp-pointed tweezers.
- Use a stainless-steel spatula to lift the entire brain from the cranial base.
- Collect the brain into a centrifuge tube (50 mL) with 20 mL of cold wash solution.
- Wash the brain twice with the cold wash solution.
4. Microdissection of the neural tissue
- Place a Petri dish on a surface surrounded by ice.
- Deposit the brain on the dish and add 20 mL of cold wash solution.
- With a scalpel, in the coronal plane, divide the brain into two blocks of tissue (rostral and caudal). As a neuroanatomical reference, perform the coronal cut at Bregma level in the anterior-posterior axis11 (Figure 1A, solid line).
- From the rostral block, extract the VZ tissue (Figure 1B), while from the caudal block remove the DG (Figure 1C).
- Dissect the VZ under a stereo microscope.
- With a Dumont forceps, hold one of the hemispheres; then, insert, at the height of the ventricle, the fine tips of a second Dumont forceps under the tissue that lines the caudate-putamen (Figure 2A).
- Open the forceps along the dorsoventral axis to separate the tissue.
- Collect the VZ tissue per individual in a centrifuge tube with 2 mL of cold wash solution. Do not pool the tissue of more than two animals.
- Repeat the microdissection in the other hemisphere.
- Store the tube containing the bilateral VZ tissue on ice and continue dissecting the DG.
- Dissect the DG from the caudal block under a stereo microscope.
- With a scalpel, make a coronal cut into the block to obtain two slices, in which the hippocampal formation is observed. As a landmark, the cut is made at -2 mm Bregma coordinates in the anterior-posterior axis according to the mouse brain atlas11 (Figure 1A, dotted line and Figure 1C).
- With a Dumont forceps, hold one of the slices, and with fine-point Dumont forceps make a horizontal cut between DG and CA1 and then perform a vertical incision between the DG and CA3 to separate the DG (Figure 2B).
- Repeat the dissection in the first slice of the other hemisphere.
- Repeat the dissection in both hemispheres in the second slice.
- Collect the four DG pieces of each vole in a centrifuge tube. Do not pool the DG tissue of more than two animals.
NOTE: If dissection of more than one animal is required, store the centrifuge tubes with the VZ or DG tissue on ice while continuing to dissect the rest of the brains. Remove all blood vessels that cover the brain tissue while dissecting. If the vessels are not discarded, the culture could be mixed with an excess of erythrocytes and disturb neurosphere formation.
5. Isolation of neural cells
- Place the centrifuge tubes inside the biosafety cabinet and wait about 10 min for the tissue fragments to precipitate by gravity.
- Remove the wash solution and add 1 mL of the warm enzymatic solution to each tube.
- Incubate the tubes at 37 °C for 10 min.
- Disintegrate the tissue fragments; pipette up and down with a 1 mL tip. Do not pipette more than 30x.
- Carry out a second incubation of 10 min at 37 °C.
- At the end of the second incubation, pipette to break up the tissues. Do not pipette more than 30x.
NOTE: After pipetting, the tissue fragments should be completely disintegrated; if they are not disintegrated, incubate for another 10 min at 37 °C and re-pipette. The digestion period should not exceed 30 min.
- Add 9 mL of N2 medium per tube to dilute the enzymatic treatment.
- Centrifuge the tubes at 200 x g for 4 min at room temperature.
- Discard the supernatant and wash with 10 mL of N2 medium.
- Centrifuge under the same conditions as step 5.8.
- Remove the supernatant from each tube and resuspend the cell pellets of the VZ and DG in 2 mL and 1 mL of the B27 medium, respectively.
- To remove any non-disintegrated tissue, filter each cellular suspension using a cell strainer (size 40 µm).
6. Neurospheres formation
- Culture the cells passed through the strainer into an ultra-low attachment, 24-well plate. Use two wells for the VZ and one well for the DG (1 mL of B27 medium/well).
- Add 20 ng/mL of FGF2 and 20 ng/mL of EGF to each well (final concentration 1x).
- Incubate at 37 °C, 5% CO2 and high humidity (90-95%). Do not disturb for 48 h (day 1 and day 2 of culture, D1-D2).
- On the third day (D3), remove half of the culture medium and replace it with fresh B27 medium (500 µL per well) supplemented with double concentration (2x) of growth factors.
- Repeat every third day, change the culture medium (half of it) and replace it with a fresh B27 medium supplemented with double concentration (2x) of growth factors.
- On days when it is not necessary to change the culture medium, add growth factors to a final concentration of 1x.
- Ensure that the neurospheres are formed around D8-D10.
- At the D10, change the complete culture medium to remove all debris.
- Collect the medium and neurospheres individually of each well in centrifuge tubes.
- Incubate for 10 min at room temperature. This procedure allows neurospheres precipitation by gravity.
- Remove the supernatant and resuspend in 1 mL of fresh B27 medium supplemented with growth factors.
- Place the neurospheres back into the same ultra-low attachment plate and incubate at 37 °C, 5% CO2.
- From D10 to D15, continue changing half of the medium and adding growth factors.
7. Passage of the neurospheres
- At D15 of the primary culture, collect the neurospheres into centrifuge tubes using 1 mL pipette. Cut the pipette tip to increase the size of the opening to avoid damage to the neurospheres.
- Incubate for 10 min at room temperature. Neurospheres precipitate by gravity.
- Remove the medium and add 1 mL of the cell detachment medium per tube.
- Incubate the tubes for 7 min at 37 °C.
- Pipette up and down with a 1 mL tip to dismantle the neurospheres.
- Dilute the cell detachment medium with 3 mL of B27 medium per tube.
- Centrifuge the cell suspension for 5 min at 200 x g.
- Discard the supernatant and resuspend each cell pellet with a fresh B27 medium supplemented with growth factors.
- Resuspend the VZ-derived cells in 4 mL of medium and the DG-derived cells in 2 mL of medium.
- Culture the cells (passage 1) in a new ultra-low attachment plate by doubling the number of wells that were used in the primary culture (4 and 2 wells for VZ and DG, respectively).
- Change half of the medium every third day and add growth factors daily.
- After 10 days (D10) in passage 1, change to adherent conditions in the next passage.
8. The passage in adherent conditions
- Before carrying out the passage 2, prepare coated plates with poly-L-ornithine and laminin.
- In 24-well plates, add 500 µL of 1x poly-L-ornithine (20 µg/mL) per well. Incubate at 37 °C overnight.
- Remove the poly-L-ornithine and wash 4x with 1x PBS (500 µL/well).
- Add 200 µL (minimum volume to cover the surface of a single well) of 1x laminin (5 µg/mL) per well and incubate for 2-3 h at 37 °C before cultivating the cells.
- Collect the neurospheres with 1 mL pipette with cut tips into a centrifuge tube.
- Incubate for 10 min at room temperature to precipitate the neurospheres by gravity.
- Discard the supernatant and resuspend the neurospheres in fresh B27 medium without growth factors.
- Aspirate the laminin from the coated plate and deposit the neurospheres into the wells using 1 mL pipettes with cut tips.
NOTE: Prevent coated wells from drying out between laminin removal and plating neurospheres.
- Divide the culture into two conditions:
- Maintain differentiated neurospheres for 6 days (D6). Change the medium every third day and add growth factors daily.
- Observe differentiation of the neurosphere-derived cells by 12 days (D12). Change the medium every third day without growth factors.
NOTE: At the end of D6 for undifferentiated or D12 for differentiation conditions, the cells can be used for conventional immunohistochemistry, cell sorting analysis, 5-Ethynyl-2´-deoxyuridine (EdU) staining, RNA extraction, among others.
Neurospheres were formed from neural stem cells isolated from the VZ and DG of both female and male adult prairie voles. About 8-10 days after starting the culture, cells should have formed the neurospheres. Note that the plate may contain debris in the primary culture (Figure 3A). However, in passage 1 the culture should only consist of neurospheres (Figure 3B).
A higher number of neurospheres were obtained from the female VZ as compared with the male VZ and DG of both females and males (Figure 4A). These data suggest that the number of neurospheres obtained depends on the proliferative zone and the vole sex. Once the neurospheres appeared (D8-D10), they were maintained for another seven days in culture, and their growth was monitored during this period. The diameter of the neurospheres was measured on D8, D11, and D14 (Table 1 and Figure 4B). The neurosphere's size (diameter) increased progressively according to the days of culture for male and female voles in both neuronal regions. Neurospheres derived from the male brains were smaller in comparison to the neurospheres derived from the female brain in both neurogenic areas (Figure 4B).
After 15 days of primary culture on floating conditions, the neurospheres were expanded in passage 1 under the same conditions. For the subsequent passage 2, the cells grew in adhesive culture, although they were able to adhere since passage 1. Adhered neurospheres were characterized at day six (D6) in the presence of growth factors (undifferentiated condition, Figure 5A) or instead until day 15 (D15) without growth factors (differentiated condition, Figure 5B).
At D6 under undifferentiation conditions, the neurosphere-derived cells expressed nestin (a marker for neural progenitors) (Figure 6). Also, it was possible to identify doublecortin (DCX) positive cells (migration cells) and the proliferation marker Ki67, which indicate the presence of either neuronal precursors or immature neurons. However, the lack of colocalization of Ki67 with DCX suggests the presence of postmitotic neuroblasts (Figure 7). Finally, at D15 under differentiation conditions, mature neurons (MAP2-positive cells) were found, as well as cells with the glial phenotype (GFAP-positive cells), which demonstrates differentiation potential of the isolated cells (Figure 8).
Figure 1: Dorsal view of an adult vole brain and its neurogenic regions. (A) The solid line at Bregma level was the anatomical reference to separate the brain into two blocks, rostral and caudal. The dotted line was the reference to divide the caudal block to obtain two slices containing the DG. (B) Coronal view of the neuronal regions exposed with the first incision, where the VZ is located. (C) Coronal view of the anatomical regions exposed with the second incision, where the DG is located. Please click here to view a larger version of this figure.
Figure 2: Anatomical references for the dissection of neurogenic regions. (A) Scheme and photograph of the coronal section from the rostral block showing the VZ location (dotted line). (B) Scheme and photograph of the coronal section from the caudal block showing the DG dissection. CPu, caudate putamen; V, ventricle; VZ, ventricular zone, DG, dentate gyrus; CA1 and CA3, regions of the hippocampus. Please click here to view a larger version of this figure.
Figure 3: Representative micrographs of neurospheres culture derived from neurogenic niches of the adult prairie vole. (A) Primary culture of neurospheres isolated from the VZ of female voles at D10. (B) Passage 1 of neurospheres derived from the VZ of female voles at D10. Scale bars = 200 µm. n= 3 for each neurogenic region and sex of the vole. Please click here to view a larger version of this figure.
Figure 4: The number and size of neurospheres depended on both sex and neurogenic source. (A) The number of neurospheres in primary culture obtained from VZ and DG in both female and male voles at D10. Data were analyzed with a one-way ANOVA followed by a Tukey’s post hoc tests. Significant differences were found between the female VZ and the rest of the groups, ***p<0.001. (B) The diameter of the neurospheres throughout D8-D14 in the primary culture depended on the vole sex. Data were analyzed with a two-way ANOVA followed by Tukey’s post hoc tests. Intra-group comparisons (differences within the same group) showed an increase in the neurosphere size between D8 vs. D11 and D14, (*p<0.05, ***p<0.001, ****p<0.0001); and D11 vs. D14 (+++ p<0.001) in the VZ and DG of female and male voles. Inter-group comparison (differences between groups in the same region) showed that female VZ and DG neurospheres are larger than male neurospheres at D11 and D14. ### p<0.0001. VZ was obtained from males and female voles (n=3, per group). 15 female and 10 male neurospheres were analyzed. DG was obtained from males and female voles (n=3, per group). 8 female neurospheres and 5 male neurospheres were processed. Please click here to view a larger version of this figure.
Figure 5: Representative images of neurospheres derived from the female VZ cultured in adhesion conditions in passage 2. (A) Neurospheres adhered in passage 2 with growth factors at D2. (B) Neurosphere-derived cells adhered in passage 2 without growth factors at D10. Scale bar = 200 µm. Please click here to view a larger version of this figure.
Figure 6: Expression of nestin in neurospheres. Representative, epifluorescence-microscopy images of nestin-positive cells derived from the VZ of both female and male adult brains at the undifferentiated stage. Scale bars = 50 µm. Please click here to view a larger version of this figure.
Figure 7: Expression of DCX and Ki67 in neurospheres. Representative epifluorescence-microscopy images of DCX-, Ki67-positive cells and merge derived from the VZ of both adult female and male brains at the undifferentiated stage. Scale bars = 25 µm. Please click here to view a larger version of this figure.
Figure 8: Expression of MAP2 and GFAP in neurospheres. Representative, epifluorescence-microscopy images of MAP2 (mature neurons) and GFAP (glial cells) positive cells derived from the VZ of both adult female and male brains at the differentiated stage. Scale bars = 50 µm. Please click here to view a larger version of this figure.
|Size of neurospheres (μm)|
|Days of culture||Sex||Mean ± SD||Days of culture||Sex||Mean ± SD|
Table 1: Quantification of the average size (diameter) of neurospheres isolated from neurogenic niches in the primary culture. Significant differences are shown in Figure 4. VZ was obtained from males and female voles (n=3, per group). Fifteen neurospheres from females and ten from males were analyzed. DG was obtained from males and female voles (n=3, per group). Eight female neurospheres and five male neurospheres were processed.
A stage to obtain a neural stem cell culture is the digestion period with the enzymatic solution, which should not exceed more than 30 min because it might decrease cell viability. The neurospheres should emerge at 8-10 days after initial culture; if they do not emerge by day 12, discard the culture and repeat the experiment, reducing the digestion period. Another issue is the blood vessels that cover the brain tissue. They should be completely removed during the dissection because the excess of erythrocytes can interfere with the neurospheres formation.
This protocol allows expanding the floating neurospheres until passage 2 and changing to adherent conditions to evaluate the neurosphere-derived cells. However, it has limitations such as a decrease in the neurogenic potential, which switches to gliogenic differentiation at successive passages as an adaptation response to in vitro conditions12. For this reason, we recommended characterizing the neurospheres in the primary culture and passage 1, and continue with the next passage only if it is required to expand the cells for experiments that do not involve elucidating differences due to the origin.
Interestingly, intrinsic differences can be found in the primary culture of neurospheres as a result of the neuroanatomical (VZ or DG) or sex-dependent (females or males) source. Thus, the number and diameter of neurospheres derived from both neurogenic regions of females are higher in comparison with males. This could be a functional difference in the female brain niche compared to that in males, which molecular mechanisms can be studied in vitro with this assay.
Cell culture of neurospheres derived from the adult brain vole is a valuable tool that could help resolve discrepancies between studies in vivo. For example, Fowler and coworkers reported that social isolation for 48 h induces an increase in 5-Bromo-2'-deoxyuridine (BrdU)-positive cells in the VZ, without affecting the DG6. In contrast, Lieberwirth et al.; demonstrated a decrease in cell proliferation in the DG13. Furthermore, in vitro culture can be a model for evaluating the molecular mechanisms in neurogenic regions that could be associated with behavioral changes in a social model such as the prairie vole. For example, it has been suggested that exposure to newborns induces, in both non-parental and parental voles, an increase of BrdU- positive cells in the DG14. The findings of this study can be confirmed using our cell culture protocol with BrdU labeling. However, although most studies on voles and other mammals use BrdU labeling to identify new cells, a disadvantage is that the labbeling might change depending on the injected doses15. EdU, another thymidine analog, is an ideal alternative to identify cells under the cell cycle phase in vitro cultures. In the same experiment, it is possible to have several periods for the incorporation of EdU, and unlike BrdU, DNA denaturation or incubation with antibodies it is not necessary for its detection. Also, EdU-positive cells can be assessed for co-localization with markers to identify the cell-division cycle (Ki67) and determine their phenotype using markers of neural stem cells or progenitors (Nestin, Sox2 and Pax6).
The neurospheres culture can be established as a model to study the effect of hormones, small molecules or drugs in the proliferation rate, neurogenesis and epigenetic modifications in the neural stem cells and progenitors of prairie voles. For example, previous studies have suggested the role of the stress hormones (like corticosterone) and estrogens in the regulation of adult neurogenesis in prairie voles, but the underlying regulatory mechanisms are unknown6.
Finally, autism spectrum disorders (ASD) and schizophrenia (SZ) are related to impairments in social cognition16,17. Interestingly, oxytocin and arginine-vasopressin have a fundamental role in social and emotional behavior, and gene expression variations in their receptors (OXTR and vasopressin 1a (V1AR), respectively) are associated with both ASD and SZ18,19,20,21. Moreover, alteration in neurogenesis and neural migration during neurodevelopment are implicated in the physiopathology of these behavioral disorders22,23,24. Thus, we propose to analyze the molecular mechanisms mediated by these hormones on neurogenesis, neural migration and other cellular events whose alterations are related to neurological disorders using prairie vole cell culture in vitro model due OXTR and V1AR receptors are found in the prairie vole hippocampus25,26.
The authors have nothing to disclose.
This research was supported by grants CONACYT 252756 and 253631; UNAM-DGAPA-PAPIIT IN202818 and IN203518; INPER 2018-1-163, and NIH P51OD11132. We thank Deisy Gasca, Carlos Lozano, Martín García, Alejandra Castilla, Nidia Hernandez, Jessica Norris and Susana Castro for their excellent technical assistance.
|Goat Anti-Mouse Alexa Fluor 488||Thermo Fisher Scientific||A-11029||RRID:AB_2534088|
|Goat Anti-Rabbit Alexa Fluor 568||Thermo Fisher Scientific||A-11036||RRID:AB_10563566|
|Goat Anti-Guinea Pig Alexa Fluor 488||Thermo Fisher Scientific||A-11073||RRID:AB_2534117|
|Antibiotic-Antimycotic||Thermo Fisher Scientific/Gibco||15240062||100X|
|B-27 supplement||Thermo Fisher Scientific/Gibco||17504044||50X|
|Collagenase, Type IV||Thermo Fisher Scientific/Gibco||17104019||Powder|
|Dispase||Thermo Fisher Scientific/Gibco||17105041||Powder|
|DMEM/F12, HEPES||Thermo Fisher Scientific/Gibco||11330032|
|Glucose||any brand||Powder, Cell Culture Grade|
|GlutaMAX||Thermo Fisher Scientific/Gibco||35050061||100X|
|HEPES||any brand||Powder, Cell Culture Grade|
|Mouse Laminin||Corning||354232||1 mg/mL|
|N-2 supplement||Thermo Fisher Scientific/Gibco||17502048||100X|
|NAHCO3||any brand||Powder, Suitable for Cell Culture|
|Neurobasal||Thermo Fisher Scientific/Gibco||21103049|
|Phosphate-Buffered Saline (PBS)||Thermo Fisher Scientific/Gibco||10010023||1X|
|Recombinant Human EGF||Peprotech||AF-100-15|
|Recombinant Human FGF-basic||Peprotech||AF-100-18B|
|StemPro Accutase Cell Dissociation Reagent||Thermo Fisher Scientific/Gibco||A1110501||100 mL|
|24-well Clear Flat Bottom Ultra-Low Attachment Multiple Well Plates||Corning/Costar||3473|
|24-well Clear TC-treated Multiple Well Plates||Corning/Costar||3526|
|40 µm Cell Strainer||Corning/Falcon||352340||Blue|
|Bottle Top Vacuum Filter, 0.22 µm pore||Corning||431118||PES membrane, 45 mm diameter neck|
|Non-Pyrogenic Sterile Centrifuge Tube||any brand||with conical bottom|
|Non-Pyrogenic sterile tips of 1,000 µl, 200 µl and 10 µl.||any brand|
|Sterile cotton gauzes|
|Sterile microcentrifuge tubes of 1.5 mL||any brand|
|Sterile serological pipettes of 5, 10 and 25 mL||any brand|
|Sterile surgical gloves||any brand|
|Syringe Filters, 0.22 µm pore||Merk Millipore||SLGPR33RB||Polyethersulfone (PES) membrane, 33 mm diameter|
|Equipment and surgical instruments|
|Biological safety cabinet|
|Motorized Pipet Filler/Dispenser|
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- McGraw, L. A., Young, L. J. The prairie vole: an emerging model organism for understanding the social brain. Trends in Neuroscience. 33, (2), 103-109 (2010).
- Fowler, C. D., Liu, Y., Ouimet, C., Wang, Z. The effects of social environment on adult neurogenesis in the female prairie vole. Journal of Neurobiology. 51, (2), 115-128 (2002).
- Yang, P., et al. Multi-omic Profiling Reveals Dynamics of the Phased Progression of Pluripotency. Cell Systems. 8, (5), 427-445 (2019).
- Reynolds, B. A., Weiss, S. Generation of neurons and astrocytes from isolated cells of the adult mammalian central nervous system. Science. 255, (5052), 1707-1710 (1992).
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- Ostenfeld, T., Svendsen, C. N. Requirement for neurogenesis to proceed through the division of neuronal progenitors following differentiation of epidermal growth factor and fibroblast growth factor-2-responsive human neural stem cells. Stem Cells. 22, (5), 798-811 (2004).
- Paxinos, G., Keith, B. J. F. The mouse brain in stereotaxic coordinates. Academic Press. (2001).
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