Stem Cell Mobilization Protocols: Filgrastim vs. Mozobil

Lily C. Trajman, Ph.D. [email protected]

Introduction

Hematopoietic stem cells (HSCs) are primitive cells capable of both self renewal and producing progenitor cells that can differentiate into all the cells of the hematopoietic system (Figure 1). Hematopoietic stem cell (HSC) transplantation is an increasingly common therapy used to treat a number of cancers,including leukemia, lymphoma, multiple myeloma, and solid tumors such as neuroblastomas and Ewing’s Sarcoma (Ali, 2015). It can also cure other types of diseases, such as sickle cell anemia, β−thalassemia, Fanconi’s Anemia and various immunodeficiency syndromes. Much research has focused on the efficient collection of hematopoietic stem cells for use in transplantation. HSCs can be found in three locations: cord blood, bone marrow (BM), and the peripheral blood (PB). Of these three options, cord blood is easily obtained but has a low total number of HSCs and its source does not allow multiple collections from the same donor. Collection of HSCs from the bone marrow is a surgical procedure requiring general anesthesia, but gives a large number of cells. Collection of PBSCs is relatively easy and repeatable, since it is not a surgical procedure, but steady state peripheral blood contains few HSCs.

There is no known cell surface protein unique to HSCs; instead, HSCs are usually identified by their elevated surface expression of the transmembrane glycoprotein CD34, which is expressed on both stem and progenitor cells (Figure 1). In bone marrow, between 1 and 3% of cells are

Figure 1: Lineage markers on Hematopoietic stem and progenitor cells. CLP: Common Lymphoid Progenitor; CMP: Common Myeloid Progenitor; MEP: megakaryocyte-erythroid prognenitor; GMP: granulocyte-macrophage progenitor.

CD34+; in peripheral blood, 0.1% of cells are CD34+ (Bender, 1994; Van Epps, 1994). A successful allogeneic transplant requires 2-5×106 CD34+ cells per kilogram of recipient body weight. Thus in order to acquire enough stem cells for an allogeneic transplant in a 70kg recipient, you would need to process between 50 and 350L of blood via apheresis to obtain a sufficient number of CD34+ cells. This is an unrealistic blood volume to process, even from multiple donors. It therefore becomes imperative to find a way to mobilize stem and progenitor cells to migrate from their niche in the bone marrow into the peripheral blood.

HSCs reside in a bone marrow microenvironment made up of stromal cells and an extracellular matrix rich in fibronectin, proteoglycans and collagen. They are retained in this microenvironment due to interactions between HSC surface proteins (notably c-kit, VLA-4, CXCR4, CD62-L and CD44) and their ligands found in the extracellular matrix of the bone marrow stroma (KL, VCAM-1, SDF-1, PSGL and HA respectively) (Nervi, 2006). Trafficking from the bone marrow requires migration through the vascular barrier into the circulatory system, and is accomplished by the downregulation and / or proteolytic cleavage of HSC cell surface receptors and their ligands. Specifically, c-kit, VLA-4, VCAM-1 and SDF-1 are subject to cleavage by proteases, while SDF-1 mRNA transcription in osteoblasts is also downregulated following G-CSF administration (Figure 2) (Nervi, 2006).

Several different growth factors, cytokines and chemotherapeutic agents have been shown to induce HSC mobilization. Thus far, three different approaches have been used with success to mobilize stem cells in humans. Researchers initially noted an increase in PBSCs during the hematopoietic recovery phase after chemotherapy, making this a viable option for HSC collection for autologous stem cell transplants (Richman, 1976). Second, hematopoietic growth

Figure 2: HSC homing in the bone marrow microenvironment. A variety of receptors expressed by HSCs bind to ligands in the extracellular matrix. Addition of G-CSF induces protease release that cleaves the extracellular portion of VCAM-1 and SDF-1 in the extracellular matrix, as well as CXCR4 and VLA-4 on the HSC. This allows trafficking of the cell out of the bone marrow. In addition, G-CSF causes downregulation of SDF-1 mRNA in osteoblasts.

factors and cytokines such as granulocyte colony stimulating factor (G-CSF), IL-7, IL-8, IL-12, and granulocyte/macrophage colony stimulating factor (GM-CSF) have been used to mobilize HSCs from the bone marrow. Third, mobilization has been prompted using Plerixafor (Mozobil), a small molecule antagonist that reversibly inhibits the interaction between CXCR4 and stromal-derivedfactor 1 (SDF-1). Blocking the binding of SDF-1 allows HSCs to exit the bone marrow and traffic to the peripheral blood. Other small molecule inhibitors have also been tested (Nervi, 2006). The first two approaches offer higher PBSC collection numbers in healthy donors, since HSCs proliferate after treatment and also migrate to the peripheral blood at a greater rate. Concurrent use of Plerixafor and G-CSF has shown synergistic mobilization of HSCs in patients with multiple myeloma and lymphoma compared to G-CSF alone (Flomenberg, 2005; Callandra, 2008).

This paper will examine the use of recombinant human G-CSF (Filgrastim (Amgen)) and Mozobil (Genzyme) for CD34+ cell mobilization in healthy donors, including the mechanism of action, side effects, and therapy potential.

Filgrastim (Amgen)

Background

The growth factor G-CSF has long been recognized as an effective method for increasing HSC proliferation and mobilization, thereby increasing the number of PBSCs collected during apheresis. It is commonly used to mobilize HSC in healthy donors. Filgrastim is a G-CSF analog produced using recombinant DNA technology. In this case, the cDNA encoding human G-CSF was inserted into the Escheria coli genome, allowing high levels of expression and easy collection from the growth medium.

Mechanism of action

Filgrastim and endogenous G-CSF function by downregulating the surface expression of SDF-1α on osteoblasts, while at the same time releasing proteases from neutrophils and monocytes that cleave the adhesion molecules Kit, VCAM1 and CXCR4 found on bone marrow HSCs. The reduction in chemokine expression by stromal cells coupled with the proteolytic cleavage of homing receptors on the HSC surface releases HSCs into the blood, with the number of CD34+ cells in the blood peaking after 4-6 days of G-CSF administration (Nervi, 2006). G-CSF also acts directly on HSCs to promote proliferation (Figure 2).

Filgrastim is administered daily for at least four days at a concentration of 10μg/kg. Early studies showed that CD34+ cells peaked after 4-6 days of administration, with higher doses producing a greater number of CD34+ cells per mL up to a plateau at 10μg/kg (Grigg, 1995). Current protocols generally “pre-treat” donors with 10μg/kg G-CSF for four days, and the first round of apheresis occurs on day five.

Dosing Schedule / Protocol

The recommended protocol for Neupogen is:

  • 10μg/kg Neupogen administered subcutaneously on days 1-5, with apheresis beginning on day 5.
  • Continued daily doses of Neupogen through day 7, to be discontinued if sufficient numbers of CD34+ cells are collected earlier.
  • The FDA data sheet can be found here:
    http://www.accessdata.fda.gov/drugsatfda_docs/label/2013/103353s5157lbl.pdf

Side Effects

Side effects related to G-CSF are generally mild to moderate and transient in nature. In a 1996 experiment involving 102 normal donors, 90% of those who received G-CSF experienced one or more side effects. The most common side effect is acute bone pain (reported by 89% of donors), followed by headache (39%), body aches (23%), fatigue (14%) and nausea and/or vomiting (12%) (Stroncek, 1996). Similarly, a larger prospective study compiled in 2009 concluded that “complete recovery is universal” after looking at adverse effects among over 2400 donors, although it also noted that transient side effects were found in 85% of donors (Pulsifer, 2009). A far more rare but serious side effect is splenic rupture following G-CSF administration, which occurs with an incidence of 1 in 5,000-10,000 donors (Akyol, 2014; Falzetti, 1999).

Other Advantages / Disadvantages

Administration of G-CSF to healthy donors has raised a number of safety concerns centered around the development of hematological malignancies following growth factor administration, especially since deregulation of genes and epigenetic changes have been noted following administration of G-CSF (Hernandez, 2007). An analysis of 2400 healthy donors found no cases of acute myelogenous leukemia or myelodysplasia at follow up appointments (median time to follow-up was 49 months); however there was one case of chronic lymphocytic leukemia and 25 reported cases of non-hematologic cancers. When compared to the incidence of various cancers in the general population there was no increase among donors who had received the G-CSF mobilization regimen (Pulsifer, 2009). This has been confirmed by various other prospective analyses in the US and Europe, although Stroncek and McCullough note that donor follow up studies must continue since malignancies induced due to G-CSF administration may not appear for 10 years or more (Stroncek, 2012).

Overall, G-CSF represents and safe and potent mechanism for mobilizing HSCs to the peripheral blood. Side effects are common but largely transient and only rarely severe enough to require hospitalization.

Mozobil (Genzyme)

Background

Plerixafor (marketed by Genzyme as Mozobil)is a small molecule inhibitor initially discovered in the context of HIV research, where it was found to inhibit HIV-1 and HIV-2 entry into cells in the 1-10nM concentration range. It is a potent and specific antagonist of the T cell co-receptor CXCR4 (used by T-lymphotrophic HIV strains to gain entry to the cell). During Phase I clinical trials an unexpected side effect of this drug was an observed increase in circulating white blood cells, specifically CD34+ hematopoietic stem and progenitor cells (DeClercq, 2009).

Mozobil can be used synergistically with Filgrastim to enhance HSC collection numbers; the combination of fouror five days of G-CSF administration plus one dose of Mozobil results in a 2.5- to 3.8-fold increase in CD34+ cells collected during one apheresis session as compared to G-CSF alone (Gazitt, 2007; Liles, 2005). It is also often used as a “rescue” drug to enhance the collection ofCD34+ cells used for autologous transplants in patients with Non-Hodgkin’s Lymphoma or Multiple Myeloma if the patient fails to mobilize sufficient numbers of cells with G-CSF alone.

Mechanism of action

Plerixafor is a small molecule CXCR4 antagonist. The interaction between CXCR4, a chemokine receptor expressed on CD34+ HSCs and HPCs in the bone marrow, and Stromal cell derived factor 1α (SDF-1α), a chemotactic cytokine produced by mesenchymal stromal cells in the bone marrow,causes CD34+ cells to be retained in the bone marrow microenvironment. Specifically, chemotaxis of CXCR4 toward SDF-1α helps HSCs home to the bone marrow; once there, the binding of CXCR4 to SDF-1α, as well as the induction of other adhesion molecules, keeps HSCs localized to the bone marrow microenvironment. Plerixafor specifically and reversibly inhibits the binding of CXCR4 to SDF-1α in a concentration dependent manner, and inhibition of this interaction is sufficient to release HSCs into circulation (Figure 3).

Plerixafor has been shown to enhance peripheral CD34+ cell numbers in healthy donors when used alone or in conjunction with a G-CSF dosing regimen. Plerixafor increases

Figure 3: Trafficking of HSCs out of the bone marrow following Plerixafor administration. Plerixafor blocks the interaction between CXCR4 on the HSC and SDF-1α in the bone marrow extracellular matrix.

peripheral CD34+ cell counts in a dose dependent manner, and the FDA has approved a dose of 0.24mg per kilogram actual weight. Following a single subcutaneous or intravenous injection of Plerixafor, an increase in circulating CD34+ cell numbers is evident within one hour, and CD34+ cell numbers peak at 9 to 10 hours post injection. Plerixafor has a half life of 3-6 hours in the plasma, and circulating CD34+ cell numbers return to baseline by 24h post-injection (Liles, 2003). If used in conjunction with G-CSF, a single dose of Plerixafor (0.24mg/kg) is administered on day four of G-CSF pretreatment, approximately 8-12 hours before apheresis begins (Steinberg, 2010). Synergystic use of Plerixafor and G-CSF increases the number of CD3+ cells per kg obtained during apheresis by 3.8-fold compared to either agent alone (Liles, 2005).

Currently Plerixafor is only approved by the FDA as a mobilizing agent when used in conjunction with G-CSF; however, initial studies indicate that Plerixafor alone can increase circulating CD34+ numbers to a similar degree as G-CSF. A subcutaneous injection of 0.24mg/kgPlerixafor results in a 10- to 15-fold increase in circulating CD34+ cells after 9 hours, comparable to the number of cells obtained after a five day dosing regimen with G-CSF (Liles, 2003).

Dosing Schedule / Protocol

The FDA-approved mobilization regimen is as follows:

  • 10μg/kg G-CSF administered in the mornings on day 1-4
  • 0.24mg/kg Mozobil administered in the evening on day 4, approximately 11 hours prior to the start of apheresis.
  • Continue Mozobil injections daily for up to 4 days.
  • The full data sheet for Mozobil can be found here:
    http://www.accessdata.fda.gov/drugsatfda_docs/label/2014/022311s015lbl.pdf

Side Effects

Most side effects were mild and transient. The most common side effects of Plerixaforinjections are diarrhea (37%) and nausea (34%), followed by injection and infusion site reactions (34%), fatigue (27%) and headache (22%). Prospective analyses indicate that Plerixafor is a safe and effective mechanism for enhancing CD34+ cell collection by apheresis (http://www.accessdata.fda.gov/drugsatfda_docs/label/2014/022311s015lbl.pdf).

Other Advantages / Disadvantages

Because Plerixafor and G-CSF function to increase the number of PBSCs via different mechanisms, there is some indication that the different mobilization regimes favor different progenitor cell populations. In macaques, dosing with Plerixafor enriches for B-, T- and Mast Cell precursors, while dosing with G-CSF enriches for for neutrophil and mononuclear phagocyte precursors. Intriguingly, dosing with both Plerixafor and G-CSF together resulted in the upregulation of a different set of genes than dosing with each individual agent.Thus not only does administration of Pleraxifor increase the number of PBSCs collected during apheresis, it may allow a more comprehensive representation of all stem and progenitor lineages found in the bone marrow (Donahue, 2009).

A comparison of mobilization withPlerixafor and G-CSF in humans has shown that mobilization with Plerixafor results in fewer CD34+ cells/kg and higher numbers of CD3+ and CD4+ cells/kg. Despite this, engraftment and reconstitution rates were comparable (Devine, 2008). Plerixafor appears to offer a faster protocol for CD34+ cell collection from the peripheral blood with fewer reported side effects than G-CSF.

Conclusions

Apheresis represents a non-invasive and easily repeatable method for collecting PBSCs from healthy donors. Because of the low level of circulating PBSCs, various methods for increasing PBSC mobilization to the blood have been tested. Among healthy donors, administration of G-CSF and Plerixafor synergistically enhances the number of PBSCs recovered following apheresis, and either agent alone produces a similar elevation in PBSC numbers and results in similar rates of engraftment. Neither G-CSF nor Plerixafor have been shown to cause serious adverse side effects in humans with any frequency. However, transient, mild side effects such as bone pain and nausea are common. Prospective studies indicate that healthy donors have universally recovered from the donation process.

References:

  • Akyol G, Pala C et al (2014). “A rare but severe complication of filgrastim in a healthy donor: splenic rupture.” Transfusion and Apheresis Science 50(1): 53-55.
  • Bender JG, Unverzagt K, Walker DE, Lee W, Smith S, Williams S, Van Epps DE (1994). “Phenotypic analysis and characterization of CD34+ cells from normal human bone marrow, cord blood, peripheral blood, and mobilized peripheral blood from patients undergoing autologous stem cell transplantation.” Clinical Immunology and Immunopathology 70: 10-18.
    Callandra G, McCarty J, McGuirk J, et al (2008). “AMD3100 plus G-CSF can successfully mobilize CD34+ cells from non-Hodgkin’s lymphoma, Hodgkin’s disease and multiple myeloma patients previously failing mobilization with chemotherapy and/or cytokine treatment: compassionate use data.” Bone Marrow Transplant 41(4):331-338.
  • DeClercq E (2009). “The AMD3100 Story: The Path to the Discovery of a Stem Cell Mobilizer (Mozobil).” Biochemical Pharmacology 77(11): 1655-1664.
  • Devine SM, Vig R, Rettig M et al (2008). “Rapid mobilization of functional donor hematopoietic cells without G-CSF using AMD3100, an antagonist of the CXCR4-SDF-1 interaction.” Blood 112: 990-998.
  • Donahue RE, Jin P, Bonifacino AC et al (2009). “Plerixafor (AMD3100) and granulocyte colony stimulating factor mobilize different CD34+ cell populations based on global gene and microRNA expression signatures.” Blood 114(12): 2530-2541.
  • Falzetti F, Aversa F, Minelli O and Tabilio A (1999). “Spontaneous rupture of spleen during peripheral blood stem-cell mobilisation in a healthy donor”. Lancet
    353(9152), 555.
  • Fiala MA, Park S et al (2016). “Remobilization of hematopoietic stem cells in healthy donors for allogeneic transplantation.” Transfusion (online preview).
  • Flomenberg N, Devine SM, Dipersio JF, et al The use of AMD3100 plus G-CSF for autologous hematopoietic progenitor cell mobilization is superior to G-CSF alone Blood 2005;106(5):1867-1874
  • Gazitt Y, Freytes CO, Akay C, et al (2007). “Improved mobilization of peripheral blood CD34+ cells and dendritic cells by AMD3100 plus granulocyte colony stimulating factor in non-hodgkin’s lymphoma patients.” Stem Cells Dev. 2007 Aug;16(4):657-66.
  • Grigg AP, Roberts AW, Raunow H et al (1995). “Optimizing dose and scheduling of Filgrastim (Granulocyte colony-stimulating factor) for mobilization and collection of peripheral blood progenitor cells in normal volunteers.” Blood 86(12): 4437-4445.
  • Hernandez JM, Castilla C, Gutierrez NC, et al. (2005). “Mobilisation with G-CSF in healthy donors promotes a high but temporal deregulation of genes.”Leukemia19:1088-1091.
  • Lack NA, Green B, Dale DC, Calandra GB, Lee H, MacFarland RT, Badel K, Liles WC, Bridger G (2005). “A pharmacokinetic-pharmacodynamic model for the mobilization of CD34+ hematopoietic progenitor cells by AMD3100.”Clin Pharmacol Ther77: 427–436.
  • Liles WC, Broxmeyer HE, Rodger E, Wood B, Hubel K, Cooper S, Hangoc G, Bridger GJ, Henson GW, Calandra G, Dale DC (2003).“Mobilization of hematopoietic progenitor cells in healthy volunteers by AMD3100, a CXCR4 antagonist.”Blood102: 2728–2730.
  • Liles WC, Rodger E, Broxmeyer HE, Dehner C, Badel K, Calandra G, Christensen J, Wood B, Price TH, Dale DC (2005). “Augmented mobilization and collection of CD34+ hematopoietic cells from normal human volunteers stimulated with granulocyte-colony-stimulating factor by single-dose administration of AMD3100, a CXCR4 antagonist.”Transfusion45: 295–300.
  • Pulsipher MA, Chitphakdithai P, Miller JP et al (2009). “Adverse events among 2408 unrelated donors of peripheral blood stem cells: results of a prospective trial from the national marrow donor program.” Blood 113(15), 3604–3361.
  • Richman CM, Weiner RS, Yankee RA (1976). “Increase in circulating stem cells following chemotherapy in man.” Blood 47(6): 1031-39.
  • Steinberg M and Silva M (2010). “Plerixafor: A Chemokine receptor-4 antagonist for mobilization of hematopoietic stem cells for transplantation after high dose therapy for non-hodgkin’s lymphoma or multiple myeloma.” Clinical Therapeutics 32(5): 821-843.
  • Stroncek DF, Clay Me, Petzoldt ML et al (1996). “Treatment of normal individuals with granulocyte-colony stimulating factor: donor experiences and the effects on peripheral blood CD345+ cell counts and on the collection of peripheral blood stem cells.” Transfusion 36(7): 601-10.
  • Stroncek DF and McCullough J (2012). “Safeguarding the Long Term Health of Hematopoietic Donors: A Continuous and Evolving Process to Maintain Donor Safety and Trust.” Expert Review of Hematology 5(1): 1-3.
  • Van Epps DE, Bender J, Lee W, Schilling M, Smith A, Smith S, Unverzagt K, Law P, Burgess J. “Harvesting, characterization, and culture of CD34+ cells from human bone marrow, peripheral blood, and cord blood.” Blood Cells 20: 411-23.
September 24, 2016

Stem Cell Mobilization Protocols: Filgrastim vs. Mozobil

Lily C. Trajman, Ph.D. [email protected] Introduction Hematopoietic stem cells (HSCs) are primitive cells capable of both self renewal and producing progenitor cells that can differentiate into all the cells of the hematopoietic system (Figure 1). Hematopoietic stem cell (HSC) transplantation is an increasingly common therapy used to treat a number of cancers,including leukemia, lymphoma, multiple myeloma, and solid tumors such as neuroblastomas and Ewing’s Sarcoma (Ali, 2015). It can also cure other types of diseases, such as sickle cell anemia, β−thalassemia, Fanconi’s Anemia and various immunodeficiency syndromes. Much research has focused on the efficient collection of hematopoietic stem cells for use in transplantation. HSCs can be found in three locations: cord blood, bone marrow (BM), and the peripheral blood (PB). Of these three options, cord blood is easily obtained but has a low total number of HSCs and its source does not allow multiple collections from the same donor. Collection of HSCs from the bone marrow is a surgical procedure requiring general anesthesia, but gives a large number of cells. Collection of PBSCs is relatively easy and repeatable, since it is not a surgical procedure, but steady state peripheral blood contains few HSCs. There is no known cell surface protein unique to HSCs; instead, HSCs are usually identified by their elevated surface expression of the transmembrane glycoprotein CD34, which is expressed on both stem and progenitor cells (Figure 1). In bone marrow, between 1 and 3% of cells are Figure 1: Lineage markers on Hematopoietic stem and progenitor cells. CLP: Common Lymphoid Progenitor; CMP: Common Myeloid Progenitor; MEP: megakaryocyte-erythroid prognenitor; GMP: granulocyte-macrophage progenitor. CD34+; in peripheral blood, 0.1% of cells are CD34+ (Bender, 1994; Van Epps, 1994). A successful allogeneic transplant requires 2-5×106 CD34+ cells per kilogram of recipient body weight. Thus in order to acquire enough stem cells for an allogeneic transplant in a 70kg recipient, you would need to process between 50 and 350L of blood via apheresis to obtain a sufficient number of CD34+ cells. This is an unrealistic blood volume to process, even from multiple donors. It therefore becomes imperative to find a way to mobilize stem and progenitor cells to migrate from their niche in the bone marrow into the peripheral blood. HSCs reside in a bone marrow microenvironment made up of stromal cells and an extracellular matrix rich in fibronectin, proteoglycans and collagen. They are retained in this microenvironment due to interactions between HSC […]
September 24, 2016

Cryopreservation of Primary Mammalian Cells

Lily C. Trajman, Ph.D. [email protected] Background Ex vivo cell-based therapies for are increasingly being used as first line treatments for a variety of diseases. However, primary cells are difficult to maintain in cell culture, and often long term culture leads to loss of pluripotency and/or functionality, rendering the cells unsuitable for use in therapy. Cryopreservation of early passage primary cells in liquid nitrogen essentially freezes the cells in time, allowing them to be used months or years later with little to no loss of functionality and pluripotency. Thus careful cryopreservation offers a better solution than continuous culture for ex-vivo cell based therapies where several doses of therapy over weeks or months are required. This paper examines cryopreservation of primary cells with a focus on leukopak and T cell protocols. Cryopreservation Protocols Cryopreservation is used for long term storage of both primary and immortalized cell lines. With the latter, optimization of freeze/thaw protocols is less critical; any surviving cells can be grown indefinitely in cell culture to achieve the required total number of cells for a given experiment. Primary cells pose a different problem. Cryopreservation is necessary because primary cells often lose their functionality or pluripotency after prolonged growth in cell culture; after thawing cryopreserved primary cells it is likewise important to minimize length of time in cell culture prior to use. Thus optimizing the cryopreservation protocol for maximum viability and retention of functionality is essential for success. A general cryopreservation protocol involves resuspending freshly isolated primary cells in a cocktail consisting on an isotonic electrolytic solution, a cryoprotectant, and a protein source, and then gradually reducing the temperature of the cells and solution to less than 150oC. Variations on media, protein source (serum or albumin) and cryoprotectant all influence the viability and functionality of the cells after thawing. In general, a cycle of cryopreservation and thawing is considered successful if cell viability is greater than 90%, cell count is within 10-15% of the number frozen, and (if applicable) cell populations are within 10% to 15% of the distribution seen in a freshly isolated sample (Ramos, 2014). Cryopreservation presents challenges at both the freezing and thawing stages. The process of freezing leaves cells susceptible to damage on two fronts: the formation of intracellular ice crystals and the effects of increased solute concentration (solution effects). Successful cryopreservation protocols balance these two factors to result in minimal cell death. The appearance of intracellular […]
September 24, 2016

Leukopak 101: A Brief Review of Apheresis

Lily C. Trajman, Ph.D. [email protected] Introduction Apheresis refers to the process by which blood is removed from a patient and separated into its constituent parts, allowing the removal of one specific component from the blood while the remainder is returned to the patient. Apheresis was first described over 100 years ago – by John Abel in 1914 – and has been used as a therapy for a number of different diseases, including sickle cell anemia and certain types of cancer (Korsack, 2016). In 1971 MD Anderson Cancer Center first used apheresis as a method for isolating peripheral blood stem cells (Korbling, 2011); subsequent breakthroughs in stem cell mobilization and cryopreservation have made apheresis the predominant method for peripheral blood stem cell collection. This paper will discuss the development of apheresis as a viable therapy as well as the current uses of apheresis in medicine. Separation protocols The main principle behind apheresis is the separation of donor blood into its component parts by either centrifugation or membrane filtration. Initial protocols relied on centrifugation to separate blood components by density, with erythrocytes at the bottom, overlaid by the Buffy Coat (comprised of granulocytes, lymphocytes, monocytes and platelets), and plasma as the top layer (Figure 1). Apheresis machines (Figure 2A) are characterized as either continuous flow (blood is withdrawn from one limb, centrifuged, and returned to the donor via another limb) or intermittent flow (blood is withdrawn, centrifuged, and the desired components returned to the patient by reversing the flow of the lumen line). Apheresis by centrifugation is the preferred method for removal of a specific cell type from the blood, and is also used for plasmapheresis (Figure 2B). Separation by membrane filtration was developed in 1980 (Sueoka, 1997) and removes small molecules from the blood based on size exclusion. Because of this, it is the favored method for extracting plasma proteins from the blood. Extracted proteins can then be passed through subsequent membranes or over an affinity column to allow removal or further concentration of the protein of interest. Separation by membrane filtration can be used to remove autoantibodies, circulating immune complexes, soluble inflammatory factors, lipids such as LDL, and paraproteins (Kaplan, 2013). A basic protocol for apheresis can be found in Appendix 1. Therapeutic Apheresis Apheresis as therapy was first used in 1952 to treat hyperviscosity due to multiple myeloma; in the 1960s it was used to treat chronic myelocytic leukemia […]
September 9, 2016

Chimeric Antigen Receptors and Personalized Immunotherapy

T cells stand at the apex of the immune surveillance system. T cells become activated after the T cell receptor (TCR) binds its cognate peptide in the context of cell surface proteins encoded by the Major Histocompatibility Complex (MHC) family of genes on an antigen presenting cell (APC). The requirement of recognizing both MHC and peptide makes TCR binding a more complex interaction than the relatively simple binding of B cell antibody to antigen. Once a TCR binds to its cognate peptide-MHC the T cell requires a second signal via costimulatory receptors in order to achieve full activation. Activated T cells then trigger cytokine release and a cascade of proliferation and differentiation begins. T cell responsiveness is thus limited in a way that B cell responsiveness is not: T cells only respond to peptides that have been processed and are presented on native MHCs, and require a second signal for full activation. Extracellular proteins, viruses or bacteria presented without processing or in the context of a foreign MHC complex do not elicit an immune response. Absence of a costimulatory signal results in T cell anergy. The ability of T cells to detect non-self antigens and initiate an immune response led researchers to investigate whether T cells were able to detect and eliminate tumor cells. Experiments with mice lacking different components of the immune system illustrated that immune surveillance does play a role in detecting and eliminating cancer (reviewed in Swann, 2007); however cancers frequently evade immune detection. Evasion is achieved via several different mechanisms. First, cancers may fail to elicit an immune response because there are no “non-self” peptides or proteins presented to T and B cells. However, cancer cells do upregulate the expression of some stress-related genes that induce a T cell response. If such a response fails to completely eliminate the malignant cells, surviving cells may then evade immune detection by downregulating the expression of MHC proteins on the surface of the cell or slowing the proteolytic process that generates peptides for display in MHC proteins. Mouse models involving cancer cells that have been engineered to present a foreign peptide or protein on the cell surface have demonstrated that the immune system is efficient at eradicating cancer cells if it can identify them as non-self (Swann, 2007). With this noted, researchers began to examine alternate ways to trigger an immune response to cancer in humans. CARs are chimeric antigen receptors that target specific (generally native) antigens. The first generation of CAR-T cells were composed of a single chain variable fragment (scFv) from an antibody […]