In Situ (In Vivo) Gene Transfer Into Murine Bone Marrow Stem Cells

Abstract

Adult bone marrow stem cell is an ideal target for gene therapy of genetic diseases, selected malignant diseases and AIDS. The in vivo approach of lentivirus vector (LV) - mediated stem cell gene transfer by intrafemoral (IF) injection can take full advantage of any source of stem cells residing in the bone cavity. Such an approach may avoid several difficulties encountered by ex vivo hematopoietic stem cell (HSC) gene transfer. We have shown that both HSC and mesenchymal stem/progenitor cells (MSC) can be genetically modified successfully by a single “in situ” IF injection in their natural “niche” in mice without any pre-conditioning. This approach may provide a novel application for treatment of human diseases, and represent an interesting new tool to study adult stem cell plasticity and the nature of unperturbed hematopoiesis.

1. Introduction

Bone marrow stem cells from adults have been viewed as the ideal target for gene- and cell-based therapy of genetic diseases, selected malignant diseases and AIDS. Under steady-state conditions, i.e. normal hematopoietic turnover and an intact bone marrow (BM) “niche”, the majority of hematopoietic stem cells (HSC) in humans cycle slowly, yet continuously [1]. In vivo vector delivery can take advantage of both the natural HSC cycling for efficient transduction, and the supportive microenvironment in bone cavity for maintaining stem cell viability and capacities. We and others have demonstrated the feasibility of in vivo gene transfer into murine adult BM HSC and/or mesenchymal stem/progenitor cells (MSC) by a single intravenous injection of a lentiviral vector (LV) [2, 3], or by a relatively localized “in situ” intrafemoral (IF) injection of a retroviral vector into mice pretreated with sublethal dose of 5-fluorouracil (5-FU) [4] or a LV into un-conditioned mice [5].

The in vivo approach of LV-mediated stem cell gene transfer may have a significant impact on disease treatment and stem cell research. First, direct gene delivery would take full advantage of any source of stem cells present in the bone cavity, including those highly “plastic” cells that can differentiate into mature, nonhematopoietic cells of multiple tissues including epithelial cells of the liver, kidney, lung, skin and GI tract [6](see review [7]). Second, this approach would avoid the difficulties encountered by ex vivo HSC gene transfer, including maintaining stem cell properties and the loss of engraftment potential. Third, it would also avoid cytokine stimulation, which may activate unwanted signaling pathways that could potentially increase the risk of non-random mutagenic events during provirus insertion. Therefore, this approach may provide a novel application for treatment of human diseases. It may also represent an interesting new tool to study adult stem cell plasticity and the nature of unperturbed hematopoiesis without the complications by in vitro manipulation or in vivo pre-conditioning.

Concentrated lentiviral vector stocks with high infectivity are essential for in vivo gene transfer. Survival surgery with intrafemoral injection is performed in mice under continuous inhalation anesthesia using aseptic techniques. Post-procedural care of animals is needed to monitor and minimize any post-injection pain. Examples of in situ gene transfer into murine HSC and MSC by IF injection are shown in Figs. 1 and 2.

FIG. 1.

GFP⁺ PBL and HSC were detected up to 4-months in primary iBM injected mice. Data shown are representative results. (A) PBL from mice 3-month post injection were stained with CD11b-APC for myeloid, and CD3e-PE (and B220-PE) for lymphoid lineages. We used 7-AAD staining to gate out dead cells. (B) Total BM cells were harvested from femur and tibia of both (for LV-CG) or injected (for LV-EMiG) hind legs 4-months post injection, and stained with Sca1-PE, c-kit-APC and lineage-markers-PerCP for HSC-enriched (Lin⁻c-kit⁻Sca1⁺) cells (i.e. R3, 0.2–0.6% of total BM). (Reproduced from ref. 5 with permission from Elsevier Science.)

FIG. 2.

GFP-expressing cells were observed in long-term culture stromal cells with varied differentiation abilities. (A) GFP⁺ adherent cells were observed in long-term stromal culture derived from iBM injected mice. Cells shown (from Inj2) have been in culture for 2 months at passage 5. (B) FACS analysis of MSC cell-surface antigens was performed after in vitro stromal culture for 7–8 passages. Dashed lines represent IgG controls. (C-F) In vitro inductive culture was conducted using MSC at 8–9 passages and stained with Alizarin Red S for osteogenic (C), with Alcian-blue for chondrogenic (D) and phase contrast microscopy or Oil Red-O for adipocytic (E and F) differentiation. (G) GFP⁺ MSCs were observed with (arrow) or without (flat arrow) osteogenic potential. (H) Representative GFP-expressing CFU-fibroblastoid unit derived from CFC-F assay using MSC-Inj2 at passage 8. Colonies with diameter more than 2mm were scored. (I) CFU-F colonies were stained with Crystal Violet and randomly collected into PCR-direct lysis buffer. Duplex real-time PCR was performed on 96-well plates, each including 2–3 no template controls and one background control derived from apparently empty space. (Reproduced from ref. 5 with permission from Elsevier Science.)

2. Materials

2.1. Concentration of VSVG-pseudotyped LV

1. Ethanol sterilizing solution (70%) is made freshly from Absolute Ethyl Alcohol (AAPER Alcohol and Chemical Co., Shelbyville, KY) and autoclaved distilled H₂O.

2. Ultracentrifuge polymer tubes for SW28 rotor (36 ml) and SW55 rotor (3.2 ml) (Beckman Instrument Inc., Palo Alto, CA).

3. X-VIVO 10 serum free medium (Lonza Baltimore Inc., Baltimore, MA).

4. Tris-buffered saline (TBS) made in distilled H₂O: 50 mM Tris (2-Carboxyethyl) phosphine^.HCl, 0.9% NaCl, and 10 mM MgCl₂. The pH is adjusted with HCl to pH 7.3. The TBS is sterilized by filtration through a 0.2 μM Corning syringe filter (Sigma-Aldrich, St. Louis, MO), and should be stored at 4 °C for several months.

5. Cryogenic vials (Fisher Scientific).

2.2. Vector preparation

1. Insulin syringe with 27-gauge needle (1/2 length) (Becton Dickinson and Company, Franklin Lakes, New Jersey).

2. Sterile Alcohol Prep Pads (Fisher HealthCare).

2.3. General anesthesia and continuous inhalation anesthesia

1. Survival IF injection is performed using sterile instruments, sterile surgical gloves, mask, cap, gown and aseptic procedures.

2. Nose-cone assembly: 20 ml syringe cover (Becton Dickinson and Company), rubber lid from Vacuum collection blood tubes, catheter and surgical tapes (Fisher Scientific).

3. Isoflurane liquid (IsoFlo, halogenated agent) is from the Abbott Laboratories (North Chicago, IL), and Buprenorphine Hydrochloride (Buprenex^R) is a Schedule III narcotic obtained through Veterinary Service.

2.4. Intrafemoral injection and injection related pain management

1. 25-gauge needle with 3-ml syringe (Becton Dickinson and Company).

2. Betadine surgical scrubs (Abbott Laboratories).

3. Electric clippers (Golden A-5 Clipper, Oster, McMinnville, TN).

3. Methods

Wild type vesicular stomatitis virus (VSV) has a broad host range extending from insects to nearly all mammals [9]. The VSV glycoprotein (VSVG) envelope also has the unique ability to withstand the shearing forces encountered during ultracentrifugation. Therefore, VSVG has been utilized to “pseudotype” gene therapy vectors such as retroviral [10], HIV-1 based [11] and FIV based [12] lentiviral vectors to broaden vector host range (tropism) and increase potency in vector stocks by concentration. In addition to the ultracentrifugation method described here, other interventions have also been reported including low-speed overnight concentration and filtration. The high infectivity in concentrated vector stocks provide the basis for efficient in vivo HSC gene transfer by IF injection.

The intrafemoral injection can be accomplished without open-wound surgery in adult mice (10 weeks and older). It can also be applied for intrafemoral cell transplantation procedure [8] with minor modification. To reduce the potential post-operational complications, a bio-bubble or biosafety cabinet is used during the operation, and the injected mice are accommodated in a specific pathogen-free facility.

The IF injection approach is limited by the applicable volume per injection and relatively low transduction frequency for clinical application. It is likely that LV-mediated stem cell gene transfer by IF injection could be further improved by pre-treatment with 5-FU or sublethal irradiation to reduce the numbers of HSC staying at G0 phase and total numbers of BM cells, by temporarily stopping blood flow (by tourniquet) or by successful repeated LV administration.

3.1. Concentration of VSVG-pseudotyped lentiviral vector

1. Vector should be kept on ice whenever it is possible during all process to reduce the loss of infectivity.

2. Sanitize rotor buckets and caps with 70% ethanol and let them air dry in a Biosafety cabinet.

3. Load the pre-sterilized ultracentrifuge tubes with vector supernatant that has been filtered through a 0.45 μM filter (see Note 1).

4. Insert tubes into rotor buckets for SW28 rotor, and put the corresponding caps on. Each pair of tube-bucket sets (number 1 with 4, 2 with 5 and 3 with 6) should be balanced, followed by tightening the cap with the alignment of the two identical numbers on cap and bucket.

5. Hook the buckets in place onto the rotor by their assigned numbers (see Note 2).

6. Position the rotor inside the Beckman L-90 Ultracentrifuge. Spin rotor counter-clockwise gently by hand to ensure a secure and correct connection.

7. Ultracentrifuge at 20,000 rpm and 4 °C for 2 hours with brake set at “low” (see Note 3).

8. Remove the tubes from buckets promptly in the hood without shaking. Discard the supernatant into bleach by aspirating pipettes carefully to avoid disturbing viral pellets.

9. Resuspend each pellet in 0.5% of starting medium volume (~180 μl) of X-VIVO 10 medium by gently shaking the tubes (inserted in a sterile 50 ml Conical tube) in a shaker for 15 minutes. Pipette up and down to fully resuspend each pellet and pool several pellets together.

10. Rinse each tubes with 300–400 μl X-VIVO 10 medium and collect them to the pooled vector tube.

11. A second ultracentrifugation is performed using a SW55 rotor at 21,000 rpm and 4 °C for 1 hour.

12. Remove the supernatant as much as possible and discard into the bleach.

13. Resuspend the vector pellets in TBS at the volume of 1:2000 of initial vector amount by gently shaking at 4 °C for 30 minutes to loose the pellet, followed by pipetting to ensure complete suspension.

14. Aliquot the concentrated LV stocks into cryogenic vials with 2–3 diluted aliquots (1:100) for titration assay and replication competent lentivirus (RCL) test (see Note 4).

15. Vector stocks should be stored at −80 °C for further usage.

3.2. Vector preparation for injection

1. At the date of IF injection, the concentrated LV stocks should be quickly thawed with cap-side up at 30°C in a water both.

2. As soon as they are thawed, mix the vector by pipetting up and down 2–3 times, and pool vector as needed, with care to avoid extensive bubbles.

3. The vector supernatant should be transferred into a 27-gauge needle insulin syringe (with 1/2 length) by pipetting into the unplugged syringe (see Note 5).

4. Plug the insert back carefully and eliminate all the air bubbles. Point the needle to an alcohol wipe during this step to collect any potential discharge.

5. The vector-containing syringe should be stored on ice during the rest of the experiment.

3.3. General anesthesia and continuous inhalation anesthesia

1. Connect the pre-sterilized sedating chamber with the outlet tubing of an anesthesia machine.

2. Complete the assembly of the nose-cone with the rubber lid on and connect it to the outlet tubing of a North American Drager NarkoMed Anesthesia System (see Note 6). Multiple sets can be connected for simultaneous operation.

3. Secure the tubing(s) on the surgery plane with heat pad to keep the animal body temperature during operation.

4. Turn on the NarkoMed Anesthesia System and set the oxygen rate at 0.6% (or 1.2% for double unites) and isoflurane at 2.5.

5. The initial general anesthesia is performed in the sedating chamber pre-filled with isoflurane at the rate of 3.

6. To control the significant post-operational pain, an analgesic agent Buprenorphine (Buprenex^R) is administered at 0.5 to 1 mg/kg by subcutaneous injection in the back of the head right after the subject showed no response to touching (see Note 7).

7. The animal is put back to the sedating chamber until the smooth/slow breathing stage is observed for deep anesthesia phase.

8. Shave the hair around the knee joints of both hind limbs with an electric clipper for a better view of the kneecaps. The left knee is prepared in case of the failure with attempts in the right knee (see Note 8).

9. The animal is transferred to surgery plane and laid on its back with nose securely inside the nose cone. Secure the nose-corn in position on the heat pad with surgical tapes.

10. Use surgical tapes to attach both forelegs by the side on the heat pad.

11. Watch the breath of the mouse and gently pinch a leg with a twister to make sure the continuous anesthesia of the subject.

3.4. Intrafemoral injection

1. These injection procedures should be practiced with a coloring solution (such as Trypan-blue) on freshly sacrificed mice.

2. Use alcohol wipes to clean the knee areas of both hind limbs.

3. Prepare a 25-gauge needle attached to a 3-mL empty syringe. Make sure to use a new needle for each knee.

4. Bend the right knee with the femur at about 75° angel with the heat pad.

5. Gently feel the kneecap with the needle to find the highest point of the femur and lodge between the condyles (see Note 9).

6. Apply gentle twisting and pressure to gain access to the intrafemoral space. Always check the insertion by turning the needle around and slightly back and forth to ensure the needle is inside the marrow cavity (see Note 10).

7. Aspirate the marrow to reduce the number of marrow cells to be transduced for higher multiplicity of infection (see Note 11).

8. Hold the femur and tibia tightly in place while retrieving the penetrating needle.

9. Re-insert the vector-containing insulin syringe needle into the marrow cavity (see Note 12). Re-check the insertion as above in step 6.

10. Inject 40–50 μl of vector supernatant slowly in 2–3 minutes with simultaneously retrieving the needle. Wait for another 2 minutes before removing the needle completely.

11. Immediately straighten the injected limb and use Betadine surgical scrubs to clean knee area (see Note 13).

12. Ear clipping to appoint identification for the injected mouse right after removing it from the nosecorn.

3.5. Post-injection pain management

1. The injected mice are allowed to emergence from general anesthesia in their cage with their mouth facing sideway (see Note 14).

2. All mice are observed for ambulation before transferring back to husbandry.

3. Twenty-four hours later, a thorough observation for behavioral signs of acute pain is performed for all injected mice, including attitude, porphyrin staining, gait and posture, and appetite (Table 1). No additional intervention is needed if the score is under 0.8.

4. For mice with score of 0.8 to 1.2, Ibuprofen is added in drinking water at 7.5 mg/kg per oral ad libitum (see Note 15). Mice with score >1.2 and obvious limping are removed from the study.

5. A repeated evaluation is conducted on Day 2 post injection.

6. A weekly weight measurement should be maintained to monitor potential chronic pain or any sign of abnormal development.

Table 1.

Post-operational evaluation criteria for pain management.

Observation	Scoring benchmark	Score
Attitude	Bright, active and alert (BAR)	0
	Burrowing/ hiding, quiet but rouses when touched	0.1
	No cage exploration when lid off, burrowing/hiding, may vocalize when head pressed, or be unusually aggressive when touched	0.4
Porphyrin staining	None	0
	Mild around eyes and/or nostrils	0.1
	Obvious on face and/or paws	0.4
Gaiting and posture	Normal	0
	Mild in-coordination when stimulated, hunched posture, mild piloerection	0.1
	Obvious ataxia or head-tilt, dragging one or both limbs, severe piloerection	0.4
Appetite	Normal, eats dry food, evidence of urine and feces	0
	No evidence of eating food, drinks but appears hydrated (by skin tent test)*	0.1
	No interest in food or treats, and/or appears dehydrated	0.4

^*The loose skin on the side, back, or shoulders is gently pulled outward to create the “tent” in the skin. If skin tent does not immediately return to position, then the mouse is dehydrated.

4. Notes

1. The tubes should be filled in full with 35–36 ml supernatant (add fresh media if needed); otherwise they may collapse during centrifugation.

2. All 6 buckets with full-filled tubes should be in place to make sure a stable ultracentrifugation even if tube(s) with water is needed as “balancer”.

3. The initiation of centrifugation should be observed to make sure that no imbalance occurs, and that the speed and vacuum reach the desired settings.

4. Prior to injection, 0.5% of the total LV production should be tested to be “free” of RCL.

5. The use of insulin syringe can minimize the loss of vector stock in the needle’s dead space.

6. For economy, the nose-cone unite can be made using a 20 ml syringe cover with both cover pieces secured by surgical tape to avoid gas leakage. A hole is generated on the longitudinal side with the size that can fit the lid of 14 ml blood collecting tubes. An open catheter needle is inserted into the syringe cover with the entrance tapped to prevent gas leakage. The catheter is connected to the outlet tubing of the Anesthesia System.

7. Surgery on the femur is painful because of large muscle mass trauma and direct bone manipulation. An average weight can be obtained from 2–3 randomly selected (male or female, separately) mice, and used as a guideline for the dose of Buprenorphine injection.

8. Hair shaving has a risk of skin rupture that increase the chance of infection. For the well being of the subject and the reduction of anesthesia time, this step can be skipped if injection on mice with hair-on can be achieved during practice.

9. A small incision can be made over the kneecap for easier observation of the condyles, especially when long-term survival of the mice is not required.

10. It is important to insert the needle with appropriate angle along with the femur. A maximum of two insertion attempts can be applied to one kneecap without serious damage to the limb.

11. The marrow aspiration step is not always successful due to the blockage of the needle during knee penetration. If the marrow aspiration is required, a second attempt can be performed with a new needle-syringe set.

12. It is essential to keep the femur and tibia in place during needle change. Make sure the vector-containing syringe is within easy reach or have a co-worker to assist on this step.

13. The execution of injection can vary from mouse to mouse. For better reference of follow-up assays, it is important to document the actual injection process, such as side of injection, number of attempts, success of marrow aspiration, and any visible leakage after removing the injecting needle.

14. Mice should regain consciousness within 5 minutes. They are generally quiet but responsive when touched.

15. Inclusion of Ibuprofen in water can lead to decreased water consumption, and should be stopped as soon as un-necessary. A close observation is needed for signs of dehydration during treatment period.