Dual Lentiviral Transduction To Generate Mutant Neural Stem Cells For Glioma Research

Summary

This protocol describes the generation of mutant murine neural stem cells (NSCs) for glioma research using sequential lentiviral transductions. Oncogenic mutations are first introduced into NSCs, followed by a bicistronic lentivirus containing reporter genes. After dual antibiotic selection, mutant NSCs enable real-time imaging of tumor growth and treatment response.

Abstract

Gliomas are the most prevalent primary brain tumors in adults, and murine modeling suggests that they may arise from oncogenic mutations in NSCs or early differentiating progenitors. Orthotopic transplantation models derived from mutant NSCs thus have the potential to yield valuable preclinical information about glioma pathogenesis, outcomes, and treatment response. These models require the introduction of an oncogenic mutation into NSCs, preferentially coupled with reporters to aid in the determination of transfection/transduction efficiency and to monitor tumor growth in vivo. This protocol outlines a dual transduction-based methodology for generating mutant NSCs suitable for in vivo tumor monitoring. In this technique, isolated NSCs are first transduced with lentiviruses that contain a gene of interest (oncogenic mutation or green fluorescent protein [GFP] control) in addition to a blasticidin-resistance element. Following blasticidin selection, cells are secondarily transduced with a bicistronic lentiviral vector expressing luciferase and red fluorescent protein (RFP), as well as a puromycin-resistance element. After puromycin selection, cells can be expanded for proliferation assays, orthotopic transplantation, or other downstream applications. This dual transduction approach allows for estimation of transduction efficiency at both infection stages as well as monitoring in vivo tumor establishment, growth, and response to therapy using noninvasive bioluminescent imaging (BLI). This system represents a useful tool for glioma research, merging the biological relevance of adult neural stem cell-derived tumor modeling with the practical benefit of real-time imaging capability, ultimately enhancing the understanding of glioma biology and the advancement of new therapeutic approaches.

Introduction

Gliomas are the most prevalent form of primary brain tumors affecting adults, constituting approximately 80% of all malignant brain neoplasms in this population¹. The majority of these are high-grade gliomas², which are typically very invasive and carry a poor prognosis^3,4 despite significant advances in neurosurgical techniques, radiation therapy, and chemotherapeutic approaches⁵. The high mortality rate in patients afflicted by these tumors highlights the need for ongoing research into glioma formation and progression.

Gliomagenesis in the central nervous system can be triggered by introducing oncogenic mutations into NSCs^6,7,8,9 or their early differentiating progeny^10,11,12 in murine modeling studies. Notably, these studies have shown that the same oncogenic mutations often fail to produce tumors when introduced into more differentiated cells¹³, establishing a cell-of-origin hierarchy where increased lineage restriction correlates with reduced susceptibility to transformation¹⁴. Genetically modified NSCs are thus commonly used to generate biologically relevant and versatile glioma models, using either transgenic techniques or orthotopic transplantation methods¹⁵. Both approaches allow for the study of de novo tumor formation in the native brain environment from an untransformed cell. Genetically engineered models allow for this study in animals with an intact immune system, often better mimicking the histology and cellular heterogeneity observed in human tumors¹⁶. However, generating these models is labor- and cost-intensive, and they frequently exhibit long latency times and low tumor formation frequencies¹⁶. In contrast, orthotopic NSC models are more readily manipulable and cost-effective. Mutant NSCs can be generated from primary or immortalized cells from either mice or humans, based on experimental needs, and are also suitable for in vitro. experimentation^16,17. Additionally, these models offer the ability to precisely adjust the developmental stage of both NSCs and recipient mice, as well as the spatial localization from which NSCs are derived and to which they are introduced. This enables researchers to investigate how age and specific brain regions influence tumorigenesis driven by particular mutations¹⁷.

This protocol outlines the methodology for generating mutant murine NSCs from primary culture, which are suitable for both in vitro and in vivo experimental applications (Figure 1). The protocol consists of two distinct transduction phases that occur in a specific temporal sequence. In the initial transduction phase, isolated murine NSCs are infected with recombinant lentiviruses carrying the target genetic construct, either an oncogenic mutation or a green fluorescent protein (GFP) control, alongside a blasticidin resistance cassette. Following this, antibiotic selection pressure is applied using blasticidin, allowing only the surviving transduced cells to proceed to a secondary transduction event. This secondary event employs a bicistronic lentiviral construct that encodes both firefly luciferase and red fluorescent protein (RFP) reporter genes, coupled with puromycin resistance elements for subsequent selection. After the selection with puromycin, the resulting dually transduced NSC populations are expanded in vitro. before subsequent experimentation. This sequential transduction approach offers several methodological advantages: (1) it facilitates a quantitative assessment of transduction efficiency at each step¹⁸, (2) it establishes stable reporter cell lines suitable for longitudinal tracking, and (3) it enables noninvasive monitoring of tumor initiation, progression kinetics, and therapeutic responses through BLI methodologies^18,19,20. It can also be used with primary or immortalized NSCs^21,22,23.

The protocol presented here addresses the pressing need for a standardized, efficient, and biologically relevant approach to generating NSC-derived glioma models from adult animals with integrated imaging capabilities. The resulting models are anticipated to yield insights that are clinically relevant regarding glioma pathogenesis, facilitate the identification of novel therapeutic targets, and present solid platforms for the preclinical evaluation of emerging treatment strategies.

Protocol

This study was conducted in accordance with the ethical standards and regulatory guidelines established by Washington University in St. Louis. For this protocol, the cortex of adult mice was micro-dissected as per the Brossier laboratory’s previously described technique²⁴ and used to isolate primary murine NSCs for subsequent expansion in non-adherent culture according to established protocol^24,25,26. All reagents used in culture are either supplied in sterile format or passed through a 0.2 µM membrane filter to ensure sterility. The reagents and the equipment used are listed in the Table of Materials.

1. Neural stem cell isolation from micro-dissected tissue

1.  Prepare Dissociation Media (DM) and Trypsin Digest Medium (TDM), as per recipes shown in Figure 2.
   1. Add 200 µL of Trypsin Digest Medium into 1.5 mL microcentrifuge tubes and keep on ice for at least 20 min prior to use. DM can be left at room temperature for later steps in the protocol.
   2. While tubes are pre-chilling, prepare base Neural Stem Cell Medium (NSC; recipe shown in Figure 2) and place NSC and Fetal Bovine Serum (FBS) at 37 °C in a water or bead bath to pre-warm. Place aliquots of supplements (B27, N2) and growth factors (EGF, FGF) in an ice bucket to thaw slowly.
      NOTE: TDM must be prepared fresh each time before use, although NSC and DM can be stored at 4 °C between uses; recipes are shown in Figure 2. Stocks of N2, B27, EGF, and FGF should be aliquoted and stored at -20 °C until use; avoid repeated freeze-thaws of reconstituted stocks (see Table of Materials for further details in preparing stocks of EGF and FGF).
2.  After TDM-containing microcentrifuge tubes are sufficiently chilled, micro-dissect tissue as previously described²⁴. After each micro-dissection, place the tissue into a microcentrifuge tube. Return the tube to ice until all dissections have been completed, as any warming of tissue may begin the enzymatic digestion process early and result in over-digestion.
3. Incubate the microcentrifuge tube(s) at 37 °C in a water or bead bath for 15 min to initiate enzymatic digestion. During incubation, prepare 10% FBS in NSC by adding 0.5 mL FBS to 4.5 mL NSC.
   1. After incubation, pipette sample(s) gently 5–10 times with a 1 mL micropipette to further dissociate cells, then neutralize trypsin by adding 200 µL of 10% FBS in NSC to each sample. Pipette up and down gently to mix.
      NOTE: It is crucial to follow the recommended time of incubation at 37 °C. Over-trypsinization may lead to cell death, while under-trypsinization can make the sample difficult to dissociate (see Table 1, Points 1–2 for this and other troubleshooting tips).
      NOTE: Either a water bath or a bead bath may be used to incubate samples; the latter may reduce the risk of contamination²⁷, but this may come with a risk of less efficient heat transfer²⁸.
4. Spin down the isolated primary cells at 630 × g for 5 min at 4 °C. Aspirate the supernatant, being careful not to disturb the cells at the bottom of the well. Wash the cells by adding 2.5 mL DM to the cell pellet and resuspending by gently pipetting up and down 3–4 times. Spin down again at 630 × g for 5 min at 4 °C, and carefully aspirate the supernatant.
5. Repeat Step 1.3 to perform a second wash. During final wash, add B27, N2, FGF, and EGF to pre-warmed base NSC medium to create complete NSC medium (recipe shown in Figure 2).
6. Aspirate the supernatant. Add 2 mL of warm complete NSC medium to each cell pellet, then resuspend by gently pipetting up and down 3–4 times. Place resuspended cells in one well of a 6-well low-attachment plate and incubate at 37 °C.
   NOTE: Although base NSC medium can be stored at 4 °C between uses, supplements and growth factors may undergo degradation at this temperature. Thus, B27, N2, FGF, and EGF should be added to NSC medium to create a complete NSC medium immediately before use.

2. Neurosphere culture establishment and expansion

1. Administer 0.5–1 mL of fresh NSC feeding medium (recipe shown in Figure 2) to each well of the NSC cultures every 3 days.
2. Monitor for neurosphere formation. When fully formed small spheres are visible, the culture is ready for passage (typically 5–10 days from initial plating).
   NOTE: Cells grow best when passaged frequently and when still in the logarithmic growth phase. Larger neurospheres are harder to dissociate and more prone to cell death during this process. Thus, earlier and more frequent passaging is recommended to promote cell health. When ready for passage, isolated neurospheres will be ~100–200 µm in diameter with tightly packed cells and smooth, well-defined borders under brightfield microscopy. Neurospheres will not display irregular projections or extensions. The center of the neurospheres will be slightly darker than the outer edges, but not excessively so; excessively dark cores often result from necrosis of central cells due to nutrient deprivation when neurospheres have been overgrown. Passaging overgrown neurospheres often results in high amounts of cell death, which can affect downstream applications. See Table 1, Points 3–4 for this and other troubleshooting tips.
3. Pre-warm the NSC medium, DM, 0.05% Trypsin-EDTA, and phosphate-buffered saline (PBS) at 37 °C using a water or bead bath. Place stocks of B27, N2, FGF, EGF, and FBS on ice until needed.
4. To passage, transfer well-grown neurospheres to a 15 mL sterile conical tube using a serologic pipette. Centrifuge at 630 × g for 1 min at 4 °C to settle down neurospheres. Aspirate supernatant, being careful to avoid disturbing the pelleted neurospheres. Wash the cells by resuspending in sterile 1x PBS, then centrifuging again at 630 × g for 5 min at 4 °C and aspirating the supernatant.
5. Add 1 mL of 0.05% Trypsin/EDTA to the cell pellet. Pipette up and down 3–5 times with a 1 mL micropipette to mix, then incubate at 37 °C in a water or bead bath for 5–6 min.
   1. After incubation, gently pipette up and down 5–10 times with a 1 mL micropipette to break up neurospheres. Neutralize trypsin by adding 1 mL 10% FBS in NSC medium; pipette gently to mix.
      NOTE: Avoid over-trypsinization and/or overly vigorous pipetting of the neurospheres, as it may damage the NSCs and affect cell viability.
6. Add 2.5 mL DM and centrifuge at 630 × g for 5 min at 4 °C. Aspirate the supernatant. Repeat the wash step with another 2.5 mL DM, followed by centrifugation and aspiration of supernatant, to remove traces of FBS in the medium.
7. Resuspend cells in 1 mL fresh complete NSC medium. Filter cell suspension by placing a 40 µm sterile filter in a 50 mL microcentrifuge tube, then adding the cell solution to the top of the filter and allowing gravity to drain the solution into the tube. Any remaining undissociated cells will be captured by the filter, while single cells will pass through to the solution below.
8. From the resuspended, filtered cells, take 10 µL for cell counting and mix with 10 µL trypan blue. Using a manual hemocytometer, load 10 µL of the sample and count viable cells (i.e., cells that are not blue) according to standard practice²⁹. Alternatively, an automated cell counter may be used, although it may be less accurate in the presence of cell debris or aggregates^30,31.
   NOTE: Generally, less than 10%–15% of the total cells should exhibit signs of cell death (e.g., unable to exclude trypan blue). Higher rates of cell death may result from over-trypsinization, excessive mechanical force in separating cells, or poor overall health of the culture prior to trypsinization (see Table 1, Points 1–3). As this can negatively affect downstream applications, it is recommended that cells be replated (Step 2.9) and expanded again (e.g., repeat Steps 2.1–2.9) if high cell death rates are observed rather than proceeding to transduction.
9. Dilute 5 x 10⁵ cells into 2 mL fresh complete NSC medium and plate into one well of an ultra-low attachment 6-well plate. Administer 0.5–1 mL of fresh NSC feeding to each well every three days to ensure optimal culture health. When fully formed, healthy spheres are visible (see Table 1, Point number 4), the culture is ready for use in Step 3 or 4.
   NOTE: Extra NSCs can be stored at this point by resuspending one million NSCs in 1 mL of serum-free cell freezing medium in a 1.5 mL cryovial and storing at -80 °C. NSCs can also be further expanded if needed by plating additional cells in extra wells of the 6-well plate or plating into T-25 flasks.

3. Blasticidin kill curve in parent cells

NOTE: Once the passaged NSCs form healthy neurospheres, cells are ready for subsequent experimentation. Prior to transduction, it is critical to determine the optimal concentration of the selection antibiotic (blasticidin, in this case). This must be performed empirically, as different populations of NSCs may have different susceptibility to the antibiotic. If this has already been determined for the given cell type, skip to Step 4.

1. Pre-warm the base NSC medium, DM, 0.05% Trypsin-EDTA, and PBS at 37 °C. Thaw stocks of B27, N2, FGF, EGF, FBS, and blasticidin. Prior to starting, or during incubation steps below, make fresh complete NSC medium and 10% FBS in NSC medium as described above.
2. Using freshly expanded neurospheres (after culture from Step 2.9 above), trypsinize and subsequently split as per steps 2.5–2.6, then resuspend in 1 mL fresh NSC for counting. Filter cells through a 40 µm sterile filter as described above to remove undissociated cells, then count cells as described in Step 2.8.
3. Resuspend 25,000 viable cells into 2.5 mL aliquots of fresh complete NSC in 15 mL conicals, with 1 aliquot prepared per antibiotic concentration to be tested. Plan to test at least 4–6 different concentrations, although more can be tested if desired.
4. Add the appropriate amount of blasticidin to each 2.5 mL aliquot to achieve the desired end concentration (see sample calculations in Table 2). Plate 500 µL of suspended cells (5,000 cells) per well in a 24-well ultra-low attachment plate, with each concentration plated in quadruplicate.
   NOTE: To achieve the correct final concentration, blasticidin stock may need to be diluted before adding to the media. Blasticidin can be diluted in NSC medium or PBS; for a 1:1000 diluted stock (10 mg/mL), add 1 mL of undiluted (10 mg/mL) blasticidin stock to 999 mL NSC or PBS. It is recommended that investigators perform all needed calculations to achieve the desired concentrations while materials are warming in Step 3.1; sample calculations shown in Table 2.
5. Incubate the cells at 37 °C. Check cells carefully using brightfield microscopy every 24 h for changes in morphology and cell/neurosphere death. Feed cells every 3 days by adding 0.5–1 mL of fresh NSC feeding medium supplemented with the correct concentration of blasticidin.
6. Determine the appropriate concentration for subsequent blasticidin selection by identifying the lowest blasticidin concentration that kills non-transduced cells between 72–120 h post-plating.
   NOTE: For adult murine NSCs, 1000 ng/mL (1 mg/mL) blasticidin was identified as the best concentration for antibiotic selection (Figure 3). Subsequent steps are performed using blasticidin at this concentration. It should be noted that the optimal blasticidin concentration can vary considerably across cell preparations, typically ranging from 0.5–10 µg/mL depending on the type of sample, passage number, and culture conditions^32,33; empirical kill curves are therefore essential rather than optional for each new cell type. See Table 1, Point number 5 for additional troubleshooting tips.

4. First lentiviral transduction

NOTE: Before proceeding with the transduction, it is essential to know the optimal multiplicity of infection (MOI) at which to transduce the lentiviral particles (LVPs). This must be determined empirically for different cell types; the methods for doing so have been previously well-described^34,35. In the depicted illustrative results, an MOI of 20 was used based on prior experience in transducing primary murine NSCs.

1. Start with expanded, untransduced NSCs forming healthy neurospheres (generated after expansion of cells in Step 2.9 above). Split, resuspend, filter, and count cells as per Steps 2.3–2.8 above, then plate cells at a density of 1.5 x 10⁶ cells per well into two wells (one for oncogenic virus, one for control) of a 6-well ultra low-adhesion plate. Incubate the plates overnight at 37 °C.
2. The following day, examine the cells. NSCs will be ready for infection if they are forming small clumps.
3. Before infecting the cells, carefully thaw the lentiviral aliquot on ice, allowing for a gradual thaw. It is critical not to rush this process.
   NOTE: For illustrative purposes, this protocol describes lentiviral transduction using two representative constructs: (1) a control vector expressing GFP under the EF1-α promoter with blasticidin resistance, and (2) an experimental vector expressing an N-terminal HA-tagged BRAF V600E oncogenic mutant under the EF1-α promoter, also with blasticidin resistance. The generation of lentiviruses is beyond the scope of this protocol but has been previously well-described^36,37.
4. After thawing, briefly warm the virus to 37 °C immediately prior to the infection. Determine the volume of LVP to add to each well, using the formula MOI x cell number/virus titer. Using a micropipette set to the calculated volume, slowly add LVPs drop-wise to the cultured cells. Swirl the plate gently to distribute, being careful not to slosh media into adjacent wells. Incubate the cells at 37 °C for 24 h.
   NOTE: Once the viral aliquot has been thawed to room temperature, it cannot be refrozen and must be utilized within 24 h. Once the aliquot has been warmed to 37 °C, it needs to be employed within a 30 min timeframe.
5. Following this incubation period, centrifuge the neurospheres at 630 × g for 5 min at 4 °C, then aspirate supernatant. Resuspend in 2 mL fresh complete NSC, without any virus or antibiotic, and plate in a fresh well of a low-attachment 6-well plate.
6. On the following day, evaluate the transduction efficiency in the GFP control cells by determining the proportion of fluorescently-tagged cells/clumps to total cells/clumps by fluorescence microscopy in multiple fields. The expected reporter expression (GFP) should be more than 80% for successful transduction.
   NOTE: If suboptimal transduction efficiency is noted, a second round of infection (repeating Steps 4.4–4.6) can be performed. If efficiency fails to improve, consider re-optimization of MOI or trialing a different preparation of lentivirus. See Table 1, Point number 6 for these and other troubleshooting tips.
7. Upon observing that the developing neurospheres appear healthy and that evidence of infection is present (GFP positive; ~80%), proceed with splitting cells per Steps 2.3–2.6 above. Resuspend the cells in fresh 5 mL of complete NSC medium containing the appropriate concentration of blasticidin (1000 ng/mL), then filter through 40 µm sterile filters to eliminate cell clumps. Count cells, then plate at approximately 5 x 10⁴ cells/mL in ultra-low-attachment T-25 flasks.
   NOTE: After initial addition of antibiotic, there may be a high rate of cell death as non-transduced cells die; this is expected, and should improve with ongoing selection.
8. Expand cells by feeding every 3 days with a feeding supplement containing blasticidin. When fully formed, healthy spheres are visible, usually 5–10 days after splitting, the culture is ready for use in Step 5 or 6.
   NOTE: After further expansion of the resistant clones, cells can be collected for Western blotting (described in Step 7) to validate that the non-fluorescently labeled, HA-tagged oncogenic construct is being appropriately expressed. If Western blotting is performed at this step, it is still recommended that expression is checked after the 2^nd transduction (in Step 7), to ensure appropriate ongoing expression.

5. Puromycin kill curve in blasticidin-resistant NSCs

NOTE: The optimal selection concentration for the second transduction must also be empirically determined. Transduced NSCs resistant to blasticidin are used at this step, and cells are maintained under blasticidin selection (1000 ng/mL) throughout, as transduction and concurrent blasticidin selection may alter sensitivity to puromycin.

1. Pre-warm the base NSC medium, DM, 0.05% Trypsin-EDTA, and PBS at 37 °C. Thaw stocks of B27, N2, FGF, EGF, FBS, blasticidin, and puromycin. Prior to starting, or during incubation steps below, make fresh complete NSC medium and 10% FBS in NSC medium as described above.
2. Using blasticidin-resistant neurospheres, trypsinize and subsequently split as per Steps 2.5–2.7, resuspending in 1 mL fresh NSC for counting and filtering to remove clumps. Count the cells as described in Step 2.8, then resuspend 25,000 viable cells into 2.5 mL aliquots of fresh complete NSC in 15 mL conicals, with 1 aliquot prepared per antibiotic concentration to be tested. Plan to test at least 4–6 different concentrations, although more can be tested if desired.
3. Add blasticidin to achieve the previously optimized concentration (1000 ng/mL = 0.25 mL of undiluted stock / 2.5 mL aliquot). Then add the appropriate amount of puromycin to each tube to achieve the desired end concentration (see sample calculations in Table 2).
4. Plate 500 µL of suspended cells (5,000 cells) per well in a 24-well ultra-low attachment plate, with each concentration plated in quadruplicate.
5. Incubate the cells at 37 °C. Check cells carefully using brightfield microscopy every 24 h for changes in morphology and cell/neurosphere death. Feed cells every 3 days by adding 0.25 mL of fresh NSC feeding medium supplemented with blasticidin and the correct concentration of puromycin.
6. Determine the appropriate concentration for subsequent puromycin selection by identifying the lowest puromycin concentration that kills cells between 72–120 h post-plating.
   NOTE: For adult murine NSCs transduced with illustrative GFP and/or BRAF-V600E lentiviruses, 100 ng/mL puromycin was empirically determined as the best concentration for antibiotic selection (Figure 4). For the example transductions, subsequent steps were performed using 1000 ng/mL blasticidin (from Step 3 above) and 100 ng/mL puromycin (from Step 5 above). See Table 1, Point number 5 for troubleshooting tips.

6. Second lentiviral transduction

NOTE: Cells remain in blasticidin-containing media during the second transduction phase to ensure that the oncogenic construct or GFP control introduced in the first transduction phase is maintained in the cells.

1. Initiate the procedure by utilizing actively proliferating, stably-transduced blasticidin-resistant NSCs (generated in Step 4). Begin by dissociating the NSCs into a single-cell suspension (Steps 2.3–2.7) to ensure uniformity during plating.
2. Plate cells at a density of 1.5 x 10⁶ cells per well in complete NSC medium with blasticidin (1000 ng/mL, as determined in Step 3) into a 6-well ultra low-adhesion plate. Incubate the plates overnight at 37 °C.
3. The following day, cells will be ready for transduction if they form small clumps. Thaw the bicistronic lentivirus (containing luciferase and RFP) on ice, then briefly warm to 37 °C immediately prior to infection. Determine the volume of LVP to add to each well, using the formula MOI x cell number/virus titer; an MOI of 20 is used here.
   1. Using a micropipette set to the calculated volume, slowly add LVPs drop-wise to the cultured cells. Swirl the plate gently to distribute, being careful not to slosh media into adjacent wells. Incubate the cells at 37 °C for 24 h.
4. The following day, centrifuge the neurospheres at 630 × g for 5 min at 4 °C and replate them in fresh complete NSC containing blasticidin (1000 ng/mL, as per Step 3).
5. The following day, evaluate the efficiency of the second transduction step by determining the proportion of RFP-positive cells/clumps to total cells/clumps by fluorescence microscopy in multiple fields, with an optimal efficiency of ~60%.
6. Upon observing that neurospheres appear healthy and that the second transduction step was effective (e.g., the majority of cell clumps are RFP+), cells are ready for dual selection. Begin by dissociating the NSCs into a single-cell suspension (Steps 2.3–2.7), then replate at approximately 5 x 10⁴ cells/mL in ultra-low-attachment T-25 flasks in complete NSC containing blasticidin (1000 ng/mL) and puromycin (100 ng/mL).
   NOTE: As in Step 4.6 above, if suboptimal transduction efficiency is noted, a second round of infection (repeating Steps 6.3–6.6) may be performed. See Table 1 for further troubleshooting tips.
7. Feed cells every 3 days with a feeding supplement containing blasticidin and puromycin.
   NOTE: Vigilant monitoring of the cultures is essential to identify dual-resistant clones, which will be characterized by their survival under both selective pressures. When fully formed, healthy spheres are visible, usually 5–10 days after splitting, the culture is ready for further validation (Step 7).

7. Validation and characterization of dually transduced NSCs

NOTE: Prior to performing further in vitro or in vivo. experimentation, further validation is recommended to ensure that HA-tagged constructs and luciferase are appropriately expressed.

1. Western blotting to validate expression of the HA-tagged construct
   1. Culture NSCs (GFP control and HA-tagged BRAF. V600E, in the provided example) in T-25 ultra-low adherence flasks until ready to split (as per Step 2.2).
      NOTE: Before proceeding to the next step, it is recommended that cultures be double-checked for ongoing GFP expression (in GFP control cells) and RFP expression (in all cells).
   2. Remove half the culture and split as per Steps 2.2–2.7, then replate in fresh complete NSC medium with blasticidin (1000 ng/mL) and puromycin (100 ng/mL) for ongoing culture expansion. Spin the remainder of the neurospheres down at 630 × g at 4 °C, then aspirate off supernatant.
   3. Subsequently, wash pelleted NSCs two times with ice-cold PBS, spinning down at 630 × g at 4 °C and aspirating off supernatant after each wash. Add approximately 200 µL of ice-cold RIPA lysis buffer supplemented with protease and phosphatase inhibitors to each sample.
   4. Allow the lysates to incubate on ice for 30 min, ensuring intermittent vortexing every 10 min. Centrifuge the samples at 14,000 × g for 15 min at 4 °C. Carefully transfer the supernatant, designated as the cleared lysate, into a fresh pre-chilled tube. Store the lysates at -80 °C in a deep freezer.
   5. Lysate can then be used for Western blotting (to validate expression of constructs) according to previously established protocols³⁸.
      NOTE: If not using immediately, store lysates at -80 °C until use. Avoid repeated freeze-thaw cycles to maintain the integrity of the sample (see Table 1, Point No. 7).
2. Bioluminescence intensity (BLI) to validate the appropriate expression of firefly luciferase
   NOTE: This protocol validates luciferase expression using BLI monitoring of live cells using specific hardware and software (see Table of Materials for further details). If using a different system for BLI detection and quantification, the methodology may vary.
   1. Split NSCs and quantify as per Steps 2.2–2.8 above, then dilute cells in complete NSC medium and plate at varying concentrations (e.g., 2.5 x 10⁴ cells, 1 x 10⁵ cells/well) in a 24-well ultra-low adherence plate (500 mL/well), with each cell concentration plated in quadruplicate. Cells are maintained in complete NSC medium containing blasticidin (1000 ng/mL) and puromycin (100 ng/mL).
      NOTE: To prevent the luciferase-secreted luminescence intensity of one well from bleeding over into adjacent wells during measurement, avoid plating NSCs in adjacent wells.
   2. Power on the imaging system at least 30 min in advance to allow the charge-coupled device camera to cool to its optimal operating temperature (approximately -90 °C). Initiate the quantification software, and sanitize the imaging chamber using 70 % ethanol, allowing it to dry completely before use.
      NOTE: Maintain the temperature of the imaging chamber at 37 °C to ensure cell viability during prolonged imaging sessions.
   3. Thaw the stock aliquots of D-luciferin, ensuring that they are shielded from light exposure. Add a volume of 5 µL of 15 mg/mL D-luciferin working solution gently to each well containing cells to achieve a final concentration of 150 mg/mL. Swirl the plate lightly to ensure even distribution of the solution.
   4. Incubate the plate at 37 °C for 10 min to facilitate the diffusion of the substrate into the cells, thereby promoting maximal light production.
   5. Position the plate within the sensitive bioluminescence imaging system. Place a grid over the plate. Select the luminescence imaging mode, commencing with auto-exposure for the exposure time. Set binning to medium, and adjust the f-stop to 1 to optimize light collection.
      1. Set the field of view to 'D' for 24-well plates, and fix the subject height at 1.5 cm. Acquire images using the imaging software. If the signal seems low, the exposure time can be manually adjusted (see Troubleshooting Table Point No. 8).
   6. Perform background correction against the control wells and quantify the relative light units as necessary for subsequent analysis using the imaging software.
      NOTE: Following validation of BLI, the cells can be orthotopically transplanted into the brains of syngeneic or immunocompromised mice, in accordance with previously established protocols¹⁹.

Results

Adult NSCs were isolated from the cortex of adult C57BL/6 mice and grown in non-adherent culture³⁹, resulting in the formation of primary neurospheres (20 – 250 µm) within 7 to 10 days (Figure 1). Cells were then expanded in vitro. Optimization of blasticidin selection was then performed in untransduced NSCs (Step 3). After 24 h of selection, concentrations at or above 1000 ng/mL (1 µg/mL) blasticidin resulted in reduced neurosphere size (

Discussion

This two-step lentiviral transduction method successfully generated genetically modified NSCs suitable for oncology research. Here, effective integration of the BRAF V600E oncogenic mutation (found in a subset of adult high-grade gliomas and used here as a proof of concept) or GFP control with dual reporter systems for bioluminescence imaging (luciferase) and fluorescence microscopy detection (RFP) was illustrated. Successive transduction at a constant MOI of 20, together with a double antibiotic selection schem...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
β-mercaptoethanol (BME)	Sigma	M3148-25ML	Use inside biological safety cabinet. Make 50 mM stocks by adding 35 mL BME to 10 mL low-glucose DMEM, sterile filter and aliquot into 500 mL aliquots for use in media preparation.
Anti-mouse IgG, HRP-linked Antibody	Cell Signal Technology	7076	Used at 1:2000 dilution in 1 % Non-fat milk
B-27 Supplement (50X),	Gibco	17504-044	At first use, thaw and make 1ml aliquots for future use; store aliquots at -20 °C. Keep supplements on ice after thawing until ready to add to media
Blasticidin S HCl (10 mg/mL)	Gibco	A1113903	Make aliquots of the 10 mg/mL stock; place in 200 mL small tubes and store at  -20 °C until use.
Bovine Serum Albumin (BSA)	Millipore-Sigma	A1470-10G	Used as 10% BSA weight/volume in 1x PBS. BSA is sticky; make sure to take care to weigh it properly and add an appropriate volume of diluent to make the exact solution.
Cell Strainer (40 µm)	Midsci	40ICS	Pre-sterilized and individually wrapped. Must be opened inside the biological safety cabinet. Do not touch with bare hands
D-Luciferin, Potassium Salt	Gold Biotechnology	eLUCK-100	Make 15 mg/mL working solution (diluted in molecular grade water) and sterile filter before use. Prepare fresh stock each time and add directly to the media to achieve final working concentration of 150 mg/mL (e.g. 5 mL working solution per 500 mL culture)
DMEM, Low-Glucose	Gibco	11885-084	Used to make base NSC medium
Epidermal Growth Factor (EGF)	Corning	354001	Supplied as a sterile lyophilized powder; to reconstitute 1000X stock, add 4.95 mL PBS and 50 µL 10% BSA (in 1x PBS) to 100 µg lyophilized EGF powder, and pipette to combine.  Make 500 µL or smaller aliquots in small tubes and store at -20° C until use for complete NSC media and NSC feeding supplement. Avoid repeat freeze-thaws.
Fibroblast Growth Factor, Basic (FGF)	R&D systems	233-FB	Supplied as a sterile lyophilized powder; to reconstitute 1000X stock, add 1.24 mL PBS and 12.5  µL 10% BSA (in 1x PBS) to 25 µg lyophilized FGF powder, and pipette to combine.  Make 500 µL or smaller aliquots in small tubes and store at -20° C until use for complete NSC media and NSC feeding supplement. Avoid repeat freeze-thaws.
Glucose, 45%	Corning	25-037-CI	Supplied as sterile solution. Do not open outside of the biological safety cabinet
HA-Tag (C29F4) Rabbit mAb	Cell Signal Technology	3724	Used at 1: 1000 dilution in 3 % Bovine serum albumin (Western blotting, to verify HA-tag expression)
HBSS, 10x	Gibco	14185-052	Supplied as sterile solution. Do not open outside of the biological safety cabinet
HEPES, 1 M	Corning	25-060-CI	Supplied as sterile solution. Do not open outside of the biological safety cabinet
IVIS 50	PerkinElmer		The instrument should be powered on 30 min before use to allow the camera to cool to optimal temperature. Living Image software (version 4.3.1) is used for quantification.
Lentivirus-Luciferase (Firefly)-2A-RFP	AMSBio	LVP324-PBS	LVP must be aliquoted and stored at -80 °C refrigerator
L-Glutamine, 200 mM	Corning	25-005-CI	Supplied as sterile solution. Do not open outside of the biological safety cabinet
Molecular-Grade Water	Corning	46-000-CM	Supplied as sterile solution. Do not open outside of the biological safety cabinet
N-2 Supplement (100x)	Gibco	17502-048	At first use, thaw and make 1ml aliquots for future use; store aliquots at -20 °C. Keep supplements on ice after thawing until ready to add to media
Nestin (E5C7I) Rabbit mAb	Cell Signal Technology	89529	Used at 1: 1000 dilution in 3 % Bovine serum albumin (Western blotting, to verify NSC marker expression)
Neurobasal Media	Gibco	21103-049	Used to make base NSC medium
Penicillin-Streptomycin (10,000 U/mL)	Gibco	15140-122	Supplied as sterile solution; aliquot and store at -20°C until use.
Phosphate Buffered Saline, 1x (PBS)	Corning	MT21040CV	Supplied as sterile solution. Do not open outside of the biological safety cabinet
Protease/Phosphatase inhibitor	Cell Signal Technology	5872S	Must be stored at -20 °C
Puromycin Dihydrochloride	Gibco	A1113803	Make aliquots of the 10mg/ml stock; place in 200 mL small tubes and store at  -20 °C until use.
RIPA Lysis Buffer, 10x	Millipore-Sigma	20-188	Store at -80 °C and make aliquots of 1x in sterile double distilled water
Serum-Free Cell Freezing Medium	Bambanker (Nippon Genetics)	CS-04-001/BB02	Dilute cells in 1ml freezing medium per cryovial to store NSCs. After addition of freezing medium to cells, cells can be placed at -80 °C for long-term storage; there is no need for slow cooling or storage in liquid N2.
Sodium Bicarbonate (7.5%)	Corning	25-035-CI	Supplied as sterile solution. Do not open outside of the biological safety cabinet
Sox2 (D9B8N) Rabbit mAb	Cell Signal Technology	23064	Used at 1: 1000 dilution in 3 % Bovine serum albumin (Western blotting, to verify NSC marker expression)
Trypan Blue	Sigma Life Science	T8154-100ML	Add 10 mL trypan blue with with 10 mL cell solution for trypan blue exclusion
Trypsin-EDTA (0.05%)	Gibco	25300-054	Aliquot and keep frozen at -20 °C until use. When thawed, it should be warmed to room temperature for 1h rather than placing in bead bath. Residual trypsin can be stored at 4 °C for one week.
Ultra-low attachment multiple well plate (6-well)	Corning	CLS3471	Supplied sterile; open inside hood only.
Ultra-low attachment multiple well plate (24-well)	Corning	CLS3473	Supplied sterile; open inside hood only.
Ultra-low attachment T25 flasks	Corning	CL4616	Supplied sterile; open inside hood only.