Summary
Orthotopic human liver metastatic uveal melanoma xenograft mouse models were created using surgical orthotopic implantation techniques with patient-derived tumor chunk and needle injection techniques with cultured human uveal melanoma cell lines.
Abstract
In recent decades, subcutaneously implanted patient-derived xenograft tumors or cultured human cell lines have been increasingly recognized as more representative models to study human cancers in immunodeficient mice than traditional established human cell lines in vitro. Recently, orthotopically implanted patient-derived tumor xenograft (PDX) models in mice have been developed to better replicate features of patient tumors. A liver orthotopic xenograft mouse model is expected to be a useful cancer research platform, providing insights into tumor biology and drug therapy. However, liver orthotopic tumor implantation is generally complicated. Here we describe our protocols for the orthotopic implantation of patient-derived liver-metastatic uveal melanoma tumors. We cultured human liver metastatic uveal melanoma cell lines into immunodeficient mice. The protocols can result in consistently high technical success rates using either a surgical orthotopic implantation technique with chunks of patient-derived uveal melanoma tumor or a needle injection technique with cultured human cell line. We also describe protocols for CT scanning to detect interior liver tumors and for re-implantation techniques using cryopreserved tumors to achieve re-engraftment. Together, these protocols provide a better platform for liver orthotopic tumor mouse models of liver metastatic uveal melanoma in translational research.
Introduction
Uveal melanoma is the most common intraocular malignant tumor among adults in the western world. During the past 50 years, the incidence of uveal melanoma (5.1 cases per million) has remained stable in the United States1,2. Uveal melanoma arises from melanocytes in the iris, ciliary body, or choroid, and it is an extremely lethal disease when it develops metastasis. The death rate of patients with uveal melanoma metastasis was 80% at 1 year and 92% at 2 years after initial diagnosis of the metastases. The time between diagnosis of metastases and death is typically short, less than 6 months, without regards to therapy3,4. The cancer spreads through the blood and tends to dominantly metastasize to the liver (89-93%)4,5. An effective mouse model is urgently needed for further investigation of liver-metastatic uveal melanoma. For translational research, there is a clear demand to generate a liver-localized metastatic uveal melanoma mouse model.
Patient-derived tumor xenograft (PDX) mouse models are expected to provide individualized medicine strategies. These models might be predictive of clinical outcomes, be useful for preclinical drug evaluation, and be used for biological studies of tumors6. Representative PDX models are ectopically tumor-implanted xenograft mice, which have tumor at subcutaneous sites. Most researchers can do surgery for subcutaneous implantation without special practice7,8. They can also monitor subcutaneous tumors easily. Although subcutaneous PDX models became popular in the research phase, they have some hurdles in moving to practical use. Subcutaneous implantation forces patient-derived tumors to engraft at a different microenvironment from the tumor origin, so that it leads to engraftment failure and slow tumor growth 9,10,11,12,13,14. Orthotopic engraftment may be a more ideal and rational approach for a PDX model because it uses the same organ as the original tumor15,16.
Recently, we developed protocols for surgical orthotopic implantation techniques of patient-derived liver-metastatic uveal melanoma tumors and needle injection techniques with a cultured human liver-metastatic uveal melanoma cell line in NOD.Cg-PrkdcscidIl2rgtm1Wjl/SzJ (NSG) mice17,18. The protocols result in consistently high technical success rates. We also established CT scanning techniques that are useful to detect interior liver tumors, and we developed re-implantation of cryopreserved tumors in the PDX platform. We found that uveal melanoma tumor xenograft models maintain the characteristics of the original patient liver tumor, including their histopathological and molecular features. Together, these techniques provide a better platform for liver orthotopic tumor models for uveal melanoma in translational research.
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Protocol
Patients enrolled in the study should provide written consent allowing the use of discarded surgical samples for research purposes and genetic studies, according to an Institutional Review Board-approved protocol. This protocol was carried out in strict accordance with the recommendations in the Guide for the Care and Use of Laboratory Animals of the National Institutes of Health and approved by the Institutional Animal Care and Use Committee (IACUC).
1. Collection of Fresh Patient-derived Tumor Tissue
- Obtain patient-derived tumor tissue from surgery or a needle biopsy in a hospital operating room.
- Put the tumor tissue in a 100 mL container containing Hanks' balanced salt solution (HBSS) solution on ice.
- Transfer the tissue into a sterile hood (Biosafety level 2) in a laboratory.
- Proceed to step 2 as soon as possible.
NOTE: For safety reasons, exclude patients with known HIV or Hepatitis B or C infections.
2. Processing of Fresh Patient-derived Tumor Tissue
- Put the tissue in a 50 mL tube containing phosphate-buffered saline (PBS) on ice. For washing the tissue, add PBS in the tube and discard PBS from the tube twice.
- Transfer the tissue into a Petri dish containing PBS on ice.
- Using sterile forceps and scissors, remove the necrotic parts of the tissue. Keep the tissue moist and cold during steps 2.3 to 2.5. For needle biopsy samples, skip step 2.3 and 2.5, and do not cut the samples.
NOTE: The necrotic tissue often breaks apart easily when touched. - Cut the tissue into 1 mm3 cubes for surgical liver implantation.
- Cut the rest of the tissue into 2 mm cubes in the Petri dish.
- Transfer them to a 1.7 mm microtube with 4% formalin for histological analysis and to another tube for genomic and proteomic analysis.
- Put the microtubes in a liquid nitrogen jar with liquid nitrogen. Transfer the tubes to a -80 °C freezer for permanent storage.
NOTE: The time between sample removal from the patient and tissue processing should not exceed 30 min.
3. Surgical Liver Implantation with Patient-derived Tumor Tissue
- Spray all objects coming into the hood for surgery with 70 % ethyl alcohol.
NOTE: This includes surgical instruments, heating pads, and anesthesia machines. - Measure the weight of a cotton swab and fabric sheet.
- Anesthetize a mouse with a 3–5% isoflurane vaporizer by placing it in the induction chamber.
- Once the mouse is fully anesthetized, place it in supine position on a heating pad. Place the isoflurane cone on the mouse's snout to inhale 1.5–3% isoflurane for maintenance of anesthesia.
NOTE: The mouse needs to be on the heating pad during the entire procedure. Lack of heating may cause hypothermia. - Confirm proper anesthesia by no reaction when the foot of the mouse is pricked with ultrafine forceps.
- Inject buprenorphine (0.6 mg/kg) subcutaneously on the flank using a 27 G needle on a micro syringe before surgery.
- Apply 70% ethyl alcohol to the abdomen and spread the fur upwards and downwards. After spreading the fur, confirm easier visualization of the skin below the left subcostal area for an easier cut. Do not shave off the fur from the abdomen.
NOTE: The fur will hide the incision site after surgery and prevent the mouse from scratching the incision post operation. However, you can shave the fur to prevent infection of the incision site according to institutional standards. - Apply iodine and let it be absorbed into the skin.
- Place a sterile surgical drape with a 2 cm hole on the mouse.
- Lift abdominal skin with curved ultrafine forceps and make a 1 cm transverse left subcostal skin incision with curved scissors.
- Insert the tip of the curved scissors beneath the skin of the incision and slightly open them to separate peritoneum from skin. Retract the scissors from the incision with closed blades.
NOTE: Opening and closing scissors inside the mouse can cause damage and bleeding. - Locate the liver under the peritoneum. Confirm a dark reddish color through the peritoneum.
- With curved scissors, make a 1 cm transverse incision in the peritoneum. If a peritoneal artery bleeds from the cutting edge, immediately stop the bleeding with cautery.
- Grab fat tissue using curved ultrafine forceps with one hand, insert the edge of a cotton swab beneath the left liver lobe and roll the swab downward with the other hand to bring out the liver.
NOTE: Grabbing fat tissue is important to keep the fat tissue from sticking to the cotton swab. - Exteriorize the liver on the cotton swab and place the liver on a non-woven absorbent fabric sheet.
NOTE: The fabric sheet plays two essential roles in stabilizing the liver and absorbing hemorrhage. - Make an incision 5 mm in width and depth using a sterile No. 11 scalpel blade to form a pocket in the parenchyma while softly pressing the incision site with the cotton swab.
- Insert the blade in parallel with the surface of the liver and cut horizontally.
- Press the incision site with the cotton swab to stop any hemorrhage.
NOTE: Do not keep the blade vertical, otherwise you will break through the liver and injure large vessels in the middle of the liver.
- Roll the cotton swab upward to open the incision site and implant a 1 mm3 cube of tumor tissue into the pocket with curved ultrafine forceps. Retract the forceps while rolling the cotton swab in reverse rotation and pressing down.
NOTE: Pressing down on the incision site with the cotton swab while retracting the forceps helps to prevent displacement of the tumor inside the pocket. - Gently take the cotton swab off the incision site after implantation. Proceed to step 3.19 as soon as possible.
- Put an absorbable hemostat on the incision site.
- Confirm hemostasis. If bleeding continues, add more hemostat on the incision site.
- Peel the liver off the fabric sheet with forceps (preferably blunt-ended) and put the liver back into the abdominal cavity.
- Suture peritoneum with double ligature using 5-0 absorbable suture.
- Suture skin with triple ligature using 5-0 absorbable suture.
NOTE: Triple ligature helps to prevent surgical incision dehiscence. - Observe the mouse until fully awake and put it back in the cage.
- Measure the weight of the cotton swab and the fabric sheet with blood for bleeding volume during the surgery. Compare them with their original weights before surgery. Reduce bleeding during the surgery to less than 10% of circulating blood volume in mouse.
4. Collecting and Processing of Cultured Human Liver Metastatic Uveal Melanoma Cell Line
- Prepare cultured cells.
- Collect cells and calculate the cell number using a cell counter.
- Prepare an appropriate amount of cell suspension for 10.0 x 106 cells in a 15 mL tube.
- Spin the tube at 300 x g for 5 min in a centrifuge at room temperature.
- Remove the supernatant in the 15 mL tube. Leave the cell pellet at the bottom of the tube.
- Add 50 μL of RPMI 1640 medium into a 1.7 mL tube.
- Cut the tip of a 200 μL tip with scissors to enlarge the tip opening.
- Add 60 μL of basement membrane matrix using a pipette with the cut tip into the 1.7 mL tube that has RPMI.
- Mix RPMI and matrix in the 1.7 mL tube. Vortex it.
- Add 110 μL of the mixture into the cell pellet in the 15 mL tube. Transfer the cell suspension into a new 1.7 mL tube.
- Keep the tube on ice before needle injection.
5. Surgical Needle Implantation of Cultured Human Liver Metastatic Uveal Melanoma Cell Line into Liver
- Follow the above protocol from steps 3.1 to 3.15.
- Collect the cell suspension with a microsyringe with a 27 G needle.
- Insert the needle along the surface of the liver and advance the tip of the needle 5 mm deeper.
- Inject 20 μL of cell suspension into the liver.
- Cauterize the insertion point of the liver to prevent the injected cells from leaking out. Confirm hemostasis.
- Follow the above protocol from steps 3.21 to 3.24.
6. CT Scan
- Place the mouse into a restrainer in the awake state.
- Wipe the tail with a sterile alcohol pad for disinfection and vasodilation.
- Inject 100 µL of CT contrast agent through the tail vein with a 27 G needle on a 1 mL syringe.
- Wait for 4 h after injection before taking the CT scan.
NOTE: It takes 4 h until the agent is taken up by liver Kupffer cells. - Four hours after injection, anesthetize the tumor-bearing mouse with 3–5% vaporized isoflurane by placing it in the induction chamber.
- Once the mouse is fully anesthetized, place it in the prone position on a CT. Place the isoflurane cone on the mouse's snout to inhale 1.5–3% isoflurane for maintenance of anesthesia.
- Confirm proper anesthesia by no reaction when the foot of the mouse is pricked with ultrafine forceps.
- Take a CT scan for 15 min.
- Ensure that the mouse until it is fully awakened after the CT scan and put it back into the cage.
- Evaluate for the existence of tumor and measure the tumor size on the CT images.
NOTE: The contrast agent enhances normal liver parenchyma so that it is easy to recognize unenhanced tumor. Do not misinterpret the gallbladder and stomach as tumor.
7. Harvesting and Processing Tissue
- Euthanize mice using CO2 followed by cervical dislocation by placing the index finger and thumb behind the skull and pulling the body by the base of the tail. Proceed to step 7.2 as soon as possible.
- Place the mouse in a supine position and spray the abdomen with 70% ethyl alcohol.
- Use sterile forceps and sterile scissors to make a 3-cm transverse incision below the xiphoid process to expose the abdominal organs.
- Excise the tumor tissue and perform steps 2.1 to 2.2.
- Cut the rest of the tumor into 2 mm cubes in the Petri dish.
- Transfer them to a cryogenic tube with cryomedium for re-implantation after cryopreservation.
- Put the tubes in a cryogenic freezing container filled with isopropanol.
- Transfer the container to a -80 °C freezer for temporary storage. Do not put the cryotubes with cryomedium directly into a liquid nitrogen tank. Freeze them slowly at a cooling rate of -1 °C/min to preserve tumor tissue.
- On the next day, transfer the tubes into a liquid nitrogen tank for permanent storage.
8. Re-implantation
- Keep tubes frozen in a liquid nitrogen jar with liquid nitrogen until ready to implant tissue. Minimize exposure of the tissue to room temperature to maintain viability and enhance chances of engraftment.
- Thaw cryopreserved tube in a 37 °C water bath.
- Perform steps 2.2–2.4.
- Implant the thawed tumor into mice as described in steps 3.1–3.24.
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Representative Results
Surgical orthotopic implantation using the liver pocket method can transplant human liver metastatic uveal melanoma tumor into the mouse liver with a high success rate of 80% within six months. The xenograft tumor engrafts in the liver as a solitary tumor without daughter nodules (Figure 1 and Figure 3A). The surgical orthotopic injection technique into the liver using microneedles successfully engrafted cultured human liver-metastatic uveal melanoma cells in the liver in all cases (Figure 2 and Figure 3B). However, some cases had dissemination around the main tumor. The contrast agent detects tumors in the liver on CT, including small tumors of 1 mm size (Figure 3B). Re-implantation of cryopreserved tumors successfully established them in the mouse liver with high success rates. The re-implanted xenograft tumors after cryopreservation retain the characteristics of the original patient tumors and pre-cryopreserved tumors.
Figure 1: Patient-derived tumor xenograft mouse model by surgical orthotopic liver implantation. Mouse was euthanized after 6 months after tumor implantation. Pigmented black tumor (black arrow) is uveal melanoma. The tumor is engrafted in the left lobe of the liver. Please click here to view a larger version of this figure.
Figure 2: Liver orthotopic human cell line-derived tumor xenograft mouse model using needle injection method. Mouse was euthanized 8 weeks after tumor injection. Pigmented black tumor (black arrow) is uveal melanoma. The tumor is engrafted in the left lobe of the liver. Please click here to view a larger version of this figure.
Figure 3: CT images of liver tumors in the left lobe of the liver. Liver tumors are detected on enhanced CT. Normal liver tissue is enhanced by contrast agent. White arrows indicate the stomach next to the liver. (A) The tumor (black arrow) that was previously shown in Figure 1. Surgical orthotopic implantation forms a solitary tumor. (B) The tumors (black arrow) shown previously in Figure 2. Needle injection method forms a cluster of many small tumors. Please click here to view a larger version of this figure.
Figure 4: Technical tips for the liver pocket method. (A-C) Left lobe (white arrows) of the liver can be exposed out of the abdomen using a cotton swab (black arrow) via a 1 cm incision. A retractor is not required to widen the incision. (D) Cotton swab presses on the incision softly. It obtains hemostasis after making an incision by the scalpel (green arrow). (E) Cotton swab rolls upward (curved red arrow). This lifts the liver parenchyma to open the incision. The tumor yellow arrow) is inserted into the liver pocket through the incision by ultrafine forceps (blue arrow). (F) Cotton swab rolls downward (curved red arrow) to prevent an inserted tumor in the pocket from backing out. Please click here to view a larger version of this figure.
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Discussion
The current orthotopic xenograft models are labor-intensive, time-consuming, and expensive to create. Orthotopic tumor xenograft mouse models for liver cancer were established more than two decades ago19,20,21. However, this technique is complicated and requires use of special equipment, such as a micro-needle holder and 6-0 to 8-0 fine sutures under a microscope. Tumor and normal liver tissue must be sewn up carefully so that the suture does not damage the fragile liver tissue. The conventional techniques lead to complications, such as hematoma and necrosis22. Recently, a modified technique was developed to solve these problems23. This modified technique uses absorbable hemostatic materials instead of suture to cover the tumor on the liver surface. However, this modified method does not completely cover the tumor within the liver parenchyma. A part of the tumor is exposed to the outside. We developed a surgical orthotopic implantation technique-the liver pocket method-to house the tumor entirely inside the parenchyma18. Our method makes a pocket in the liver to provide a natural environment for tumors. The liver pocket method is simpler than the conventional technique, allowing us to finish implantation into the liver within a few minutes from the beginning of the operation. This method results in formation of a solitary tumor in the liver and does not trigger metastases, at least for as long as we observed the mice, whereas needle injection of a single cell suspension tends to disseminate as intra-hepatic metastases17. A solitary tumor is more appropriate to evaluate tumor growth and would be useful to assess efficacy in a drug trial.
Compared to the original liver pocket method18, we have modified our methods to enhance techniques of implantation. First, a retractor was not used during surgery to minimize the size of the incision. When the incision is smaller, we can shorten sewing time in surgery. With a 1 cm incision in the abdomen, we can easily bring the left lobe outside with a cotton swab (Figure 4A-C). Second, a cotton swab plays three important roles by stopping hemostasis after making the liver pocket, opening the liver pocket to be able to insert a tumor chunk and retaining the tumor chunk in the pocket without pushing the tumor back (Figure 4D-F). Average bleeding volume was approximately less than 10% of circulating blood volume in mice. Less bleeding provided great confidence in surgery. Third, a fabric sheet is useful for fixing the liver lobe outside the abdomen. The liver lobe sticks to the sheet and thus it prevents the lobe from sliding back into the abdomen (Figure 4C). One can easily cut the liver surface with a scalpel and inject a needle to the liver surface. As a result, fragile liver tissue is not injured.
We present have two troubleshooting tips for this method. First, when a small incision site is used, sometimes a left lobe is not visible. In this situation, the left lobe is likely sticking to the diaphragm. Insert blunted-edge forceps between the left lobe and the diaphragm to peel the lobe off. Second, when a tumor chunk is placed in the liver pocket with forceps, the tumor can stick to the forceps and pull back with it. Press the incision with a cotton swab while retreating the forceps. This works well to prevent the dislocation of the tumor out of the pocket.
Xenograft tumors are surrounded by mouse tissue, even though they are orthotopically implanted. Human stromal cells in patient-derived tumors are inevitably replaced by mouse stromal cells. Ideally, the mouse model had better provide human stromal tissue around tumors. Chimeric humanized liver mouse or humanized immune mouse models would be helpful to study the engraftment of uveal melanoma and to evaluate whether the drug metabolism is the same as a human-liver or human-immune environment24,25.
Orthotopic liver tumor xenograft mouse models require verification of tumor establishment with imaging studies. The commercially available CT contrast agent, developed for mouse liver CT images, allows detection of interior liver tumors in the live state on CT. The contrast agent specifically enhances normal liver on the CT. It is easy to distinguish the unenhanced site of the tumor26. The agent detects tiny tumors less than 1 mm (daughter nodules) around main tumors on CT. The agent can be tolerated by the mouse, and makes it possible to monitor liver tumors periodically. The agent would be used to evaluate efficacy of anti-cancer drugs against liver-localized xenograft tumors.
Generally, it is recommended to maintain PDX models at a relatively low passage number (less than 10) to conserve genetic and histological integrity of the original patient-derived tumor27,28,29. Most researchers refrain from making multiple passages of the PDX models to reduce the number of passages and animals. Once patient-derived tumors are temporarily preserved in a freezer, we are able to control PDX models at a lower passage number without wasting mice. This is called biobanking strategy. A cancer biobank is a rational approach to maintain tumor characteristics and to reduce the number of mice28,30. Establishing a proper biobanking method can adjust the supply of PDX models to meet the patient's treatment plan or a mouse drug efficacy trial in the future. We achieved re-implantation of cryopreserved tumors for cancer biobanking. We hope that this success facilitates PDX platform use in the near future.
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Disclosures
The authors have nothing to disclose.
Acknowledgments
We are thankful to M. Ohara, K. Saito, and M. Terai, for reviewing the manuscript. The authors acknowledge critical review for editorial and English assistance of this manuscript by Dr. R. Sato at Fox Chase Cancer Center. The work described herein was supported by the Bonnie Kroll Research Fund, the Mark Weinzierl Research Fund, the Eye Melanoma Research Fund at Thomas Jefferson University, The Osaka Community Foundation, and JSPS KAKENHI Grant Number JP 18K15596 at Osaka City University. Studies in Dr. A. Aplin's laboratory were supported by NIH grant R01 GM067893. This project was also funded by a Dean's Transformative Science Award, a Thomas Jefferson University Programmatic Initiative Award.
Materials
| Name | Company | Catalog Number | Comments |
| Materials, tissues and animals | |||
| Buprenorphine | |||
| CO2 tank | |||
| Cryomedium | |||
| Exitron nano 12000 (Alkaline earth metal-based nanoparticle contrast agent) | Miltenyl Biotec | 130-095-700 | |
| HBSS 1x, with calcium & magnesium | Corning | 21-020-CM | |
| Human liver metastatic uveal melanoma cell line | |||
| Human uveal melanoma tissue in the liver | All tissue handling should be done in a Biosafety Level 2 hood. Be careful when working with human tissue; always use gloves and avoid direct skin contact. Assume patients may have been infected with HIV or other highly transmissible organisms. Do not process samples known to carry infections. | ||
| Iodine | |||
| Isoflurane | Purdue Products | 67618-150-17 | |
| Isopropanol | Fisher scientific | A416-1 | Avoid direct contact to skin and eye and inhalation of anesthetic agent. |
| Liquid nitrogen | |||
| Matrigel HC | BD | 354248 | |
| NOD.Cg-PrkdcscidIl2rgtm1Wjl/SzJ (NSG) mice | Jackson Lab | 5557 | 4 to 8 weeks old |
| PBS 1x, without calcium and magnesium | Corning | 21-031-CM | |
| RPMI 1640 | Corning | 10-013-CV | |
| Sterile alcohol prep pad (70% isopropyl alcohol) | Nice-Pak products | B603 | |
| 4% paraformaldehyde phosphate buffer solution | Wako | 163-20145 | |
| 70% Ethyl alcohol solution | Fisher Scientific | 04-355-122 | |
| Name | Company | Catalog Number | Comments |
| Equipments | |||
| Absorbable hemostat | Johnson and Johnson | 63713-0019-61 | |
| Autoclave | |||
| Body weight measure | |||
| Cautery | Bovie Medical | MC-23009 | |
| Cell counter | |||
| Centrifuzer | |||
| Cotton swab | |||
| Cryo freezing container | NALGENE | 5100-0001 | |
| Cryotube | SARSTEDT | 72.379 | |
| Curved scissors | World Precision Instruments | 503247 | |
| Curved ultrafine forceps | World Precision Instruments | 501302 | |
| Fabric sheet | |||
| Freezer | |||
| F/AIR Filter Canister | Harvard Apparatus | 600979 | |
| Heating pad | |||
| Isoflurane vaporizer | Artisan Scientific | 66317-1 | |
| Liquid nitrogen | |||
| Liquid nitrogen jar | Thermo Fisher Scientific | 2123 | |
| Micro-CT scan | Siemens | ||
| Needle holder | World Precision Instruments | 501246 | |
| Petri dishes | Fisher Scientific | FB0875713 | |
| Pipette | |||
| Spray bottle | |||
| Sterile hood | Biosafety level 2 cabinet | ||
| Sterile No.11 scalpel | AD Surgical | A300-11-0 | |
| Straight forceps | World Precision Instruments | 14226 | |
| Surgical drape | |||
| Tail vein restrainer | Braintree Scientific | TV-150-STD | |
| Water bath | |||
| 1 mL TB syringe with 27 G needle | BD | 309623 | |
| 1.7 mL tube | Bioexpress | C-3260-1 | |
| 5-0 PDO Suture | AD Surgical | S-D518R13 | |
| 15 mL conical tubes | AZER SCIENTIFIC | ES-9152N | |
| 27 G needle | BD | 780301 | |
| 27 G needle | Hamilton | 7803-01 | |
| 50 mL conical tubes | AZER SCIENTIFIC | ES-9502N | |
| 50 µL micro syringe | BD | 80630 | |
| 50 µL micro syringe | Hamilton | 7655-01 | |
| 100 mL container | Fisher Scientific | 12594997 | |
| 200μL tip |
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