Bone metastasis models do not develop metastasis uniformly or with a 100% incidence. Direct intra-osseous tumor cell injection can result in embolization of the lung. We present our technique modeling primary bone tumors and bone metastasis using solid tumor graft implantation into bone, leading to reproducible engraftment and growth.
Primary bone tumors or bone metastasis from solid tumors result in painful osteolytic, osteoblastic, or mixed osteolytic/osteoblastic lesions. These lesions compromise bone structure, increase the risk of pathologic fracture, and leave patients with limited treatment options. Primary bone tumors metastasize to distant organs, with some types capable of spreading to other skeletal sites. However, recent evidence suggests that with many solid tumors, cancer cells that have spread to bone may be the primary source of cells that ultimately metastasize to other organ systems. Most syngeneic or xenograft mouse models of primary bone tumors involve intra-osseous (orthotopic) injection of tumor cell suspensions. Some animal models of skeletal metastasis from solid tumors also depend on direct bone injection, while others attempt to recapitulate additional steps of the bone metastatic cascade by injecting cells intravascularly or into the organ of the primary tumor. However, none of these models develop bone metastasis reliably or with an incidence of 100%. In addition, direct intra-osseous injection of tumor cells has been shown to be associated with potential tumor embolization of the lung. These embolic tumor cells engraft but do not recapitulate the metastatic cascade. We reported a mouse model of osteosarcoma in which fresh or cryopreserved tumor fragments (consisting of tumor cells plus stroma) are implanted directly into the proximal tibia using a minimally invasive surgical technique. These animals developed reproducible engraftment, growth, and, over time, osteolysis and lung metastasis. This technique has the versatility to be used to model solid tumor bone metastasis and can readily employ grafts consisting of one or multiple cell types, genetically-modified cells, patient-derived xenografts, and/or labeled cells that can be tracked by optical or advanced imaging. Here, we demonstrate this technique, modeling primary bone tumors and bone metastasis using solid tumor graft implantation into bone.
Mouse models of human and animal disease are becoming increasingly popular in biomedical research. The utility of using mice in this context is that their anatomy and physiology are very similar to humans. They have a relatively short gestation period and time in post-natal life to achieve maturity, and are largely associated with a relatively low cost and ease of housing, albeit increasing costs of development or purchase are associated with greater degrees of genetic modification, immunodeficiency, and/or humanization1. Use of inbred strains results in a largely uniform animal population prior to study inclusion. A complete knowledge of their genome suggests a high degree of similarity to humans. Orthologous molecular targets for many disease processes have been identified in the mouse genome and there is now an extensive library of mouse-specific reagents that are easily obtainable. Therefore, they provide the opportunity for relatively high-throughput analysis in a more rapid and less expensive manner when compared to larger animal models1. In addition, with the advent of genetic editing strategies that allows for the overexpression or deletion of certain genes either globally or in a cell type specific manner and/or constitutively or in an inducible manner, they represent a very biologically useful model system for the investigation of human and animal diseases2.
Cancer is one field in which mouse models have great utility. Genetic mouse models of cancer rely on modulation of the expression of either oncogenes or tumor suppressor genes, alone or in combination, for cells to undergo oncogenic transformation. The injection of primary or established tumor cell lines into mice is also performed. The introduction of either cell lines or tissues from humans or other animal species, including mice, remains the most widely used model of cancer in vivo. The use of cells and tissues from dissimilar species (xenografts) in immunocompromised mice is most commonly performed2. However, the use of allograft tumor cells or tissues where both the host and recipient are of the same species allows for the interaction with an intact immune system when combined with the same host mouse strain in syngeneic systems3.
Primary bone tumors or bone metastasis from solid tumors result in painful osteolytic, osteoblastic, or mixed osteolytic/osteoblastic lesions3,4. These tumors compromise bone structure, increasing the risk of pathologic fracture, and leave patients with limited treatment options. Primary bone tumors metastasize to distant organs, with some types capable of spreading to other skeletal sites. In breast cancer patients, bone is the most common site of first metastasis and the most frequent first site of presentation of metastatic disease5,6. In addition, disseminated tumor cells (DTCs) are present in the bone marrow prior to the diagnosis of, and predict the development of, metastasis in other organs7. Therefore, it is believed that cancer cells present in bone are the source of cells that ultimately metastasize to other organ systems. Many mouse models of solid tumor metastasis exist that develop metastasis predominantly in the lung and lymph nodes, and depending on the tumor type and injection technique, potentially other organ systems3. However, mouse models of bone metastasis are lacking that dependably, reproducibly produce site specific skeletal metastasis and develop bone metastasis before mice reach early removal criteria from primary tumor burden or metastasis to other organs. We have reported a model of the primary bone tumor osteosarcoma that relies on the surgical implantation of a solid tumor allograft into the proximal tibia of mice8. Bone tumors formed in 100% of mice and 88% developed pulmonary metastasis. This incidence of metastasis exceeds what is commonly reported clinically in people (~20-50%), but is of great interest since the lung is the most common site of metastasis for osteosarcoma9,10,11. While this model is advantageous in modeling primary bone tumors, it also has great utility in modeling bone metastasis from other osteotropic solid tumors such as breast, lung, prostate, thyroid, hepatic, renal, and gastrointestinal tumors.
The rationale for the development of this model was to develop an alternative to the traditional intra-osseous injection typically into the proximal tibia or distal femur to model primary bone tumors or bone metastasis12. Our primary goal was to alleviate a known limitation of this technique i.e., tumor embolization of the lung. This results in the engraftment of these embolic tumor cells and “artifactual metastasis” that do not recapitulate the complete metastatic cascade from an established primary bone tumor that metastasizes to the lungs8,13. This would also be the situation when an established bone metastasis spreads to a distant site. In addition, this technique was, also, developed to produce a model of bone metastasis that would ensure a greater incidence of engraftment and growth of tumors in bone and at a uniform site when compared with orthotopic or intravascular injection techniques. This model has distinct advantages over these described techniques. This model involves controlled, consistent delivery of tumor cells into the bone. It, also, avoids artifactual lung metastasis following pulmonary embolization and establishes a baseline uniform study population. There is the benefit of site-specific tumors with this model without the risk of early removal criteria resulting from primary tumors or metastasis to other organs. Lastly, this model has great utility for modification, including the use of patient-derived xenografts.
The model presented has similarities to direct cell suspension injection into bone following a surgical approach followed by either injection through the cortex or delivery into the marrow cavity after making a small defect in the cortex (with or without reaming out the medullary cavity)8,14,15,16,17. However, the implantation of a tumor allograft makes this technique distinctly different. Therefore, the purpose of this report was to demonstrate this model of primary bone tumors and bone metastasis from solid tumors, which overcomes many limitations of previously described models. Research groups with experience in cell culture, mouse models, mouse anesthesia and surgery, and mouse anatomy are well equipped to reproduce our technique to model primary bone tumors or bone metastasis in mice.
All described animal experiments were approved by the institutional animal care and use committee of University of Cambridge, Cambridge, UK.
1. Preparation of cell lines
2. Animals
3. Subcutaneous tumors
4. Surgical implantation of subcutaneous tumor fragments
5. Serial and end point assessment
A positive result would be associated with tumor engraftment and progressive tumor growth over time. Depending on the tumor type, intraosseous tumor growth may be associated with progressive hind limb lameness, but many tumors do not cause lameness despite signs of attendant bone disease. Successful engraftment was documented with advanced imaging, whereby there would be progressive radiographic, µCT, or µMRI changes in the proximal tibia associated with the bone phenotype of the cell line of interest (osteolytic, osteoblastic, or mixed osteolytic/osteoblastic metastasis) (Figure 1)8. In our previous report, the cortical defect created for tumor graft implantation was visible in the proximal tibia 1 week after implantation (Figure 1A). By week 2, there was visible osteolysis and bone remodeling adjacent to the cortical defect. From weeks 2-5, there was progressive bone destruction and the formation of new bone associated with tumor engraftment and growth (Figure 1B-E). For cell lines with reporter genes, bone changes would be accompanied by increases in fluorescence or bioluminescence imaging outputs over time. Acute lameness may be an indicator of impending or actual pathological bone fracture, necessitating immediate and careful radiographic evaluation of the mouse, and possibly early removal from the study. Depending on the tumor cell line being studied, bone destruction may occur rapidly, well ahead of metastasis. This is of specific importance in studies involving primary bone tumors, since mice may have to be removed from the study prior to the development of clinically-relevant metastasis, namely to the lung. In these cases, surgical amputation of the tumor-bearing limb is recommended to permit the study of minimal residual disease and subsequent metastasis when using primary bone tumors, as we have reported previously4,8. While complete hind limb amputation in people is not typically performed, this is standard of care in veterinary medicine, of which the similarities in the clinical and molecular characteristics of human and canine osteosarcoma are well-documented. Hind limb amputation in mice is, therefore, relevant for many animal cell lines. In addition, amputation is a viable treatment method in mice since limb-sparing procedures used in people are not achievable in small rodents such as mice. In mice bearing primary bone tumors, inappetence, progressive weight loss, general poor body condition and difficulty breathing (increased rate or effort) may signal the development of tumor metastasis to the lung and other organs. Confirmation by bioluminescent imaging, µCT, or µMRI imaging is recommended to confirm metastasis, and affected animals should be euthanized.
A negative result, most likely resulting from the lack of tumor engraftment, should be suspected if there is no evidence of progressive changes (osteolytic, osteoblastic, or mixed osteolytic/osteoblastic lesions, depending on the cell line being studied) on radiography or µCT examination of the implanted tibia. For cell lines carrying reporter genes, the lack of an increase in fluorescence or bioluminescence signaling over time by optical imaging would also support a conclusion that the tumor has failed to engraft. Lack of or poor engraftment could be associated with surgical site infection. However, this would exhibit specific clinical signs such as redness, swelling, or discharge most commonly in the early post-operative period (first 1-2 weeks after surgery). Animals could potentially also be febrile and display a lack of activity or hunched posture in the early post-operative period due to surgical site infection. Harvesting and maintaining the allografts under sterile conditions during preparation and proper sterile preparation of the limb, sterile intraoperative technique, and complete wound closure will minimize the likelihood of a post-operative surgical wound complication resulting in infection and failure of tumor engraftment.
In our previous report8, when including both the reported pilot and definitive studies, 16 out of 16 mice (100%) implanted with osteosarcoma fragments progressed to osteolytic bone lesions and 14 out of these 16 mice (88%) developed metastasis. All metastases were observed to the lung and diagnosed by histology as early as 3 weeks post-implantation8. Additional animals necropsied at 1 (n=2) and 2 (n=2) weeks after implantation did not reveal any evidence of lung metastasis.
Figure 1: Serial radiography. Serial radiography (in order) performed weekly at (A) 1, (B) 2, (C) 3, (D) 4, and (E) 5 weeks after intratibial implantation of a subcutaneous tumor allograft using a primary bone tumor cell line (osteosarcoma). Over time, there was progressive bone destruction and the formation of new bone associated with tumor engraftment and growth. This figure has been modified with permission from ref8. Please click here to view a larger version of this figure.
This report documents our model to create primary bone tumors or bone metastasis following the intratibial implantation of a tumor allograft. We believe that there are several critical steps in this process. A safe anesthetic plane should be established for both subcutaneous injection of the tumor cell suspension and intratibial placement of the resultant tumor fragments. There should be sterile preparation of the surgical site for both removal of the subcutaneous allograft and intratibial placement of the allograft. Tumor allograft fragments should be created uniformly for subsequent intratibial implantation. An appropriately sized defect in the proximal tibia should be created for implantation of tumor fragments. The surgical skin wound should be closed completely to reduce the risk of post-operative wound complications. The last critical step is that if extracellular matrix is used as part of the tumor cell suspension for subcutaneous injection, the use of manufacturer recommended handling instructions should be done to avoid solidification of the suspension prior to injection. Proper sterile preparation of the limb, sterile intraoperative technique, and complete wound closure will minimize the likelihood of a post-operative surgical wound complication.
Features of the implanted tumor and the surgical technique are critical for successful and reproducible results for incorporation into your study population. Initially, it should be verified that the tumor xenograft/allograft is implanted directly into the medullary cavity of the tibia, making sure that the tumor tissue is fully inside the bone. Modifications to prevent or limit displacement of the graft outside of the bone would initially be placement of bone wax or bone cement in the bone defect or either gel foam or a subcutaneous fat graft over the hole in the bone. An alternative would be to insert the tumor allograft through a hole on the lateral aspect of the proximal tibia, created under the cranial tibial muscle. The presence of the cranial tibial muscle would then serve as a biological cover over the bone defect, limiting potential displacement of the tumor allograft. However, this is a more aggressive surgical approach with the potential for neuromuscular trauma on the lateral aspect of the proximal tibia. Care should be taken to make sure that the tumor allografts are created to be of uniform size prior to implantation. That being said, tumor allografts should not exceed 1 mm in any dimension, since larger tumor sizes in any biological system are less likely to engraft than smaller grafts when there is not a direct, immediate vascular supply provided to the graft at the time of implantation. We have reported successful results with a final implanted tumor volume of 0.5 mm3 but anticipate combined tumor fragment volumes up to 1 mm3 will achieve successful results. However, the maximum implantable tumor fragment volume has not been determined. As compared with cryopreserved tissue, fresh tumor xenografts and allografts are more likely to survive, although we have not observed a problem with the practice of using carefully cryopreserved tumor allografts. Snap-frozen allografts should not be implanted. In addition, dissection of the capsule and outer portion off of the subcutaneous tumor will prevent any contribution from this layer of the tumor, which is largely fibrous and vascular with a significant immune infiltrate rather than consisting largely of tumor cells proper. An alternative method to creating tumor allografts from the larger tumor mass directly with a scalpel blade is by using a small biopsy needle that creates a cylindrical tumor “core” that can then be cut to a predefined length prior to implantation. Use of a biopsy needle or core biopsy punch may be beneficial in creating uniform tumor fragments for implantation, for after two uniform dimensions (width and depth) have been created by the biopsy process, they can then be cut to an appropriate length prior to implantation. If reporter genes such as luciferase or fluorescent proteins are being used in the implanted tumor cells, evaluation of tumor fragments at the time of implantation or 1 week after implantation will indicate relative contribution from tumor cells in the implanted tumor fragments, as well as viability.
As with any procedure, there are limitations in addition to our reported benefits of this technique. Metastasis is a multi-step process which requires a cell from its primary location to successfully execute invasion into and migration through the extracellular matrix, intravasate the tumor’s blood supply, survive in circulation until it arrives to its final metastatic destination, extravasate into the organ proper and then either stay in a dormant state or proliferate to create a metastatic lesion, and modification of the normal tissue architecture to become a clinically detectable metastasis18. Currently used mouse models of bone metastasis rely largely on either direct orthotopic injection of tumor cells into the normal site of the primary tumor, intravascular injection into the left ventricle of the heart or peripherally into the tail veins (lateral or dorsal caudal veins), or direct intra-osseous injection of the tumor cell suspension3. However, recent reports trying to improve bone metastasis models have advocated injection into the illiac, caudal, or femoral arteries19,20,21. All of these methods incorporate one or more steps within the bone metastatic cascade, except direct intra-osseous injection or in our case implantation of a tumor allograft, which only models end tissue modification. Therefore, this is one inherent limitation of our technique. An additional limitation is that as currently described, it requires propagating the allografts in an intermediate mouse host, which requires usage of additional mice and results in the implantation of a tumor that is not 100% pure tumor cells. The allograft will possess some stromal contribution provided by tumors growing in a subcutaneous location. A potential alternative to minimize this subcutaneous stromal contribution would be the injection of cells subcutaneously in a biological substrate such as an extracellular matrix or scaffold. Alternatively, orthotopic injection into the primary site of the tumor, followed by tumor removal and graft preparation could be performed.
The method outlined in this report has key advantages when compared to existing methods of modeling primary bone tumors or bone metastasis following the direct intra-osseous injection of tumor cells. As discussed, we and others have observed direct embolization of the lungs and occasional immediate death following intratibial injection of a cell suspension8,13. This results from a pressurized injection of tumor cells into the marrow cavity, after which tumor cells enter the peripheral blood supply through bone marrow vascular sinusoids and are carried to the lung by venous return. These events are comparable mechanistically to that observed with pulmonary or venous thromboembolism associated with pressurized placement of implants into the marrow cavity associated with total joint arthroplasty in people and animals. We and others hypothesize that this embolic phenomenon to the lungs may be cell-line specific (unpublished observations). However, very detailed steps have been reported to limit the creation of these embolic artifactual metastases12. Initially, there should be adequate trypsinization and creation of a uniform single cell suspension for injection, which should be stored on ice to prevent cell clumping. The needle should be properly placed through the proximal tibial growth plate, after which small injection volumes (~10 µL) and resistance-free injections into the marrow cavity should be performed. Despite this, tumor cell delivery to the lungs, and also to other organs, can be readily observed by bioluminescence imaging following intra-osseous and intravascular injection techniques, suggesting that it is possible that cells generating this bioluminescent signal in the lungs and other organs could give rise to distant metastasis (unpublished observations)3. We have found that another benefit of this technique is that a single tumor grows in a uniform bone site within the study population, reducing animal-to-animal variability. This enables the establishment of a uniform study population and ready comparison between treatment groups. This is in contrast to bone metastasis that develop following orthotopic or intravascular injection, where it is possible for cells to metastasize to different skeletal sites and grow with different kinetics, making comparisons between study groups challenging. Tumor growth in one anatomical site following implantation of our tumor allografts also avoids the possibility of mice being removed due to metastasis to other organ systems or primary tumor burden, as can be seen with intravascular and orthotopic injection techniques. In addition, we have observed a 100% engraftment following tumor allograft implantation, which supports a uniform study population and avoids unnecessary use of additional mice to achieve appropriate sample size numbers. This also overcomes the limitation of incomplete and inaccurate injection following percutaneous injection of tumor cell suspensions into bone.
This technique has a large degree of versatility which has resulted in modifications that are currently being used in our research and has the potential for great utility for future applications. Initially, while we employ the proximal tibia as our intra-osseous implantation site, other common sites of primary bone tumors or bone metastasis can readily be used, including but not limited to the distal femur and proximal humerus. However, the surgical approach to these sites as well as to areas such as the pelvis and spine require a gradually more extensive surgical approach when compared to the proximal tibia. In addition, sites such as the pelvis, spine, skull, radius, and ulna are not benefited by adequate bone stock for tumor implantation. For bone-invasive malignances such as with oral tumors, placement of the tumor allografts adjacent to bone or within the oropharynx may also recapitulate the progression of invasive tumors3,22. In vitro and in vivo experiments often employ co-culture or co-injection techniques to determine the effects of different cell types on a cell type of interest. Therefore, the potential exists for injection of two or more cell types simultaneously to create the subcutaneous tumors, which can then be implanted into bone using this technique. Genetically modified cells can be used alone or in combination with other normal or genetically modified cells to create the tumor allografts. With the growing interest in precision medicine and creating in vitro and in vivo models of human cancer, patient-derived xenograft transplantation into mice is gaining popularity. Recently, a model of breast cancer bone metastasis has been described where patient-derived xenografts implanted orthotopically metastasized to human bone discs implanted subcutaneously in nude mice23. From this, patient-derived xenografts could be directly implanted into bone using our model, without the need for an intermediate host for subcutaneous propagation. This would allow for the rapid evaluation of tumor kinetics and ability to evaluate multiple treatment combinations potentially from just a single patient biopsy, not requiring a large tumor mass for allograft preparation. The increasing use of organoids in cancer research could also be employed in our model24. This would require needle or pipette injection of these cell aggregates through the hole in the tibial cortex. We would, therefore, recommend sealing of the cortical defect (as discussed above) to limit the possibility of organoid migration outside of the bone and marrow cavity. As we have performed, cells can be labelled and tracked by optical (bioluminescence or fluorescence) or advanced (digital radiography, µCT, or µMRI) imaging allowing for the assessment of tumor growth over time. This also has the benefit of minimizing animal numbers since the same animals are imaged over time.
In summary, this report demonstrates our technique modeling primary bone tumors and bone metastasis using solid tumor graft implantation into bone.
The authors have nothing to disclose.
The authors acknowledge the critical contribution of Dr. Beth Chaffee, DVM, PhD, DACVP to the development of this technique.
#15 scalpel blade | Henry Schein Ltd. | 75614 | None |
6-well tissue culture plates | Thermo Fisher Scientific | 10578911 | Used for mincing tumor pieces. Can also be used for cell culture |
Abrams osteosarcoma cell line | Not applicable | Not applicable | None |
Anesthesia machine with isoflurane vaporiser and oxygen tank(s) | VetEquip | 901805 | None |
Animal weighing scale | Kent Scientific | SCL- 1015 | None |
BALB/c nude mouse (nu/nu) | Charles River Ltd. | NA | 6-8 weeks of age. Male or female mice |
Bone cement | Depuy Synthes | 160504 | Optional use instead of bone wax |
Bone wax | Ethicon | W31G | Optional |
Buprenorphine | Animalcare Ltd. | N/A | Buprecare 0.3 mg/ml Solution for Injection for Dogs and Cats |
Carbon dioxide euthanasia station | N/A | N/A | Should be provided within animal facility |
Cell culture incubator set at 37 °C and 5% carbon dioxide | Heraeus | Various | None |
Chlorhexidine surgical scrub | Vetoquinol | 411412 | None |
Cryovials (2 ml) | Thermo Scientific Nalgene | 5000-0020 | Optional if cryopreserving tumor fragments |
D-luciferin (Firefly), potassium salt | Perkin Elmer | 122799 | Optional if cell line of interest has a bioluminescent reporter gene |
Digital caliper | Mitutoyo | 500-181-30 | Can be manual |
Digital microradiography cabinet | Faxitron Bioptics, LLC | MX-20 | Optional to evaluate bone response to tumor growth |
Dimethyl sulfoxide (DMSO) | Sigma Aldrich | 1371171000 | Optional if cryopreserving tumor fragments |
Dulbecco’s modified Eagle’s medium | Thermo Fisher Scientific | 11965092 | None |
Ethanol (70%) | Sigma Aldrich | 2483 | None |
Fetal bovine serum | Thermo Fisher Scientific | 26140079 | None |
Forceps, Dumont | Fine Science Tools, Inc. | 11200-33 | None |
Freezer (– 80 °C) | Sanyo | MDF-794C | Optional if cryopreserving or snap freezing tumor fragments |
Hemocytometer | Thermo Fisher Scientific | 11704939 | Can also use automated cell counter, if available |
Hypodermic needles (27 gauge) | Henry Schein Ltd. | DIS55510 | May also use 25G (DIS55509) and 30G (Catalog DIS599) needles |
Ice | N/A | N/A | Ideally small pieces in a container for syringe and cell suspension storage |
Iris scissors | Fine Science Tools, Inc. | 14084-08 | None |
Isoflurane | Henry Schein Ltd. | 1182098 | None |
IVIS Lumina III bioluminescence/fluorescence imaging system | Perkin Elmer | CLS136334 | Optional if cell line of interest has bioluminescent or fluorescent reporter genes |
L-glutamine | Thermofisher scientifc | 25030081 | None |
Liquid nitrogen | British Oxygen Corporation | NA | Optional if cryopreserving or snap freezing tumor fragments |
Liquid nitrogen dewar, 5 litres | Thermo Fisher Scientific | TY509X1 | Optional if cryopreserving tumor fragments |
Matrigel® Matrix GFR, LDEV-Free, 5 ml | Corning Life Sciences | 356230 | Optional. Also available in 10 ml size (354230) |
Microcentrifuge | Thermo Fisher Scientific | 75002549 | Pellet cells at 1200 rpm for 5-6 minutes |
Mr. Frosty freezing containiner | Fisher Scientific | 10110051 | Optional if cryopreserving tumor fragments |
NAIR Hair remover lotion/oil | Thermo Fisher Scientific | NC0132811 | Can alternatively use an electric clipper with fine blade |
Penicillin/streptomycin | Sigma-Aldrich | P4333 | None |
Scalpel handle, #7 Short | Fine Science Tools, Inc. | 10007-12 | User preference as long as it accepts #15 scalpel blade |
Small animal heated pad | VetTech | HE006 | None |
Stereomicroscope | GT Vision Ltd. | H600BV1 | None |
Sterile phosphate-buffered saline (PBS) | Thermo Fisher Scientific | 10010023 | Use for injections and also as part of the surgical scrub, alternating with chlorhexidine |
Tissue adhesive (sterile) | 3M Corporation | 84-1469SB | Can alternatively use non-absorbable skin suture (6-0 size) |
Trypan blue | Thermo Fisher Scientific | 5250061 | None |
Trypsin-EDTA | Thermo Fisher Scientific | 25300054 | Use 0.05%-0.25% |
Tuberculin syringe (1 ml with 0.1 ml gradations) | Becton Dickinson | 309659 | Slip tip preferred over Luer |
Vented tissue culture flasks, T-75 | Corning Life Sciences | CLS3290 | Can also use smaller or larger flasks, as needed |