Cancer Chemotherapy & Biotherapy: Principles & Practices, 4th Edition


Brian P. Monahan

Carmen J. Allegra

The folate-dependent enzymes represent attractive targets for antitumor chemotherapy because of their critical role in the synthesis of the nucleotide precursors of DNA (Fig. 6.1). In 1948, Farber and associates1 were the first to show that aminopterin, a four-amino analog of folic acid, could inhibit the proliferation of leukemic cells and produce remissions in acute leukemia cases. Their findings ushered in the era of antimetabolite chemotherapy and generated great interest in the antifolate class of agents. Since then, the clinical value of antifolate compounds has been proven in the treatment of a variety of hematologic and nonhematologic malignancies. Their clinical application has also extended to the treatment of non-neoplastic disorders, including rheumatoid arthritis,2 psoriasis,3 bacterial and plasmodia infections,4 and opportunistic infections.5 The antifolates are one of the best understood and most versatile of all the cancer chemotherapeutic drug classes (Table 6.1).


Substitution of an amino group for the hydroxyl at position 4 of the pteridine ring is the critical change in the structure of antifolate compounds that leads to their antitumor activity. This change transforms the molecule from a substrate to a tight-binding inhibitor of dihydrofolate reductase (DHFR), a key enzyme in intracellular folate homeostasis. The critical importance of DHFR stems from the fact that folic acid compounds are active as coenzymes only in their fully reduced tetrahydrofolate form. Two specific tetrahydrofolates play essential roles as one-carbon carriers in the synthesis of DNA precursors. The cofactor 10-formyltetrahydrofolate provides its one-carbon group for the de novo synthesis of purines in reactions mediated by glycineamide ribonucleotide (GAR) transformylase and aminoimidazole carboxamide ribonucleotide (AICAR) transformylase. A second cofactor, 5,10-methylenetetrahydrofolate (CH2-FH4), donates its one-carbon group to the reductive methylation reaction, converting deoxyuridylate to thymidylate (Fig. 6.1). In addition to contributing a one-carbon group, 5,10-methylenetetrahydrofolate is oxidized to dihydrofolate, which must then be reduced to tetrahydrofolate by the enzyme DHFR for it to rejoin the pool of active reduced-folate cofactors. In actively proliferating tumor cells, inhibition of DHFR by methotrexate (MTX) or other 2,4-diamino antifolates leads to an accumulation of folates in the inactive dihydrofolate form, with variable depletion of reduced folates.6, 7, 8, 9, 10, 11, 12 Folate depletion, however, does not fully account for the metabolic inhibition associated with antifolate treatment because the critical reduced-folate pools may be relatively preserved even in the presence of cytotoxic concentrations of MTX. Additional factors may contribute to MTX-associated cytotoxicity, including metabolism of the parent compound to polyglutamated derivatives and the accumulation of dihydrofolate polyglutamates as a consequence of DHFR inhibition.6, 7, 13, 14, 15 Methotrexate and dihydrofolate polyglutamates represent potent direct inhibitors of the folate-dependent enzymes of thymidylate and purine biosynthesis.16, 17, 18, 19, 20, 21Thus, inhibition of DNA biosynthesis by 2,4-diamino folates is a multifactorial process consisting of both partial depletion of reduced-folate substrates and direct inhibition of folate-dependent enzymes. The relative roles of each of these mechanisms in determining antifolate-associated metabolic inhibition may depend on specific cellular factors that vary among different cancer cell lines and tumors.


Various heterocyclic compounds with the 2,4-diamino configuration have antifolate activity and include pyrimidine analogs such as pyrimethamine and trimethoprim 18, 19, 20, 21, 22, 23, 24, 25; classical pteridines such as aminopterin and MTX2; and compounds with replacement of the nitrogen at either the 5 or 8 position, or both, with a carbon atom, such as the quinazolines (trimetrexate, piritrexim) 22, 23 and 10-ethyl-10-deazaaminopterin (10-EDAM, Edatrexate).24 Investigators have designed antifolate analogs directed at targets other than DHFR, including those folate-dependent enzymes required for the de novo synthesis of purines and thymidylate synthase. A host of potent thymidylate synthase (TS) inhibitors such as 10-propargyl-5,8-dideazafolate (PDDF, CB3717)26 and closely related compounds raltitrexed (ZD1694, Tomudex) and ZD9331,27 pemetrexed (LY231514, Alimta),28 1843U89,29 and 5,8-dideazatetrahydrofolic acid (DDATHF, lometrexol) and LY231514,30 both inhibitors of GAR transformylase, have been investigated.

Figure 6.1 Sites of action of methotrexate (MTX), its polyglutamated metabolites (MTX[(Glun]), and folate byproducts of the inhibition of dihydrofolate reductase, including dihydrofolate (FH2) and 10-formyldihydrofolate (10-CHO-FH2). Also shown are 5,10-methylenetetrahydrofolate (CH2-FH4), the folate cofactor required for thymidylate synthesis, and 10-formyltetrahydrofolate (10-CHO-FH4), the required intermediate in the synthesis of purine precursors. AICAR; aminoimidazole carboxamide ribonucleotide; AMP, adenosine monophosphate; dUMP, deoxyuridylate; dTMP, thymidylate; GAR; glycineamide ribonucleotide; GMP, guanosine monophosphate; IMP, inosine monophosphate. (From DeVita VT, Hellman S, Rosenberg SA, eds. Cancer: Principles and Practice of Oncology. Philadelphia: JB Lippincott, 1989:349–397.)


Mechanism of action

Inhibition of dihydrofolate reductase leads to partial depletion of reduced folates
Polyglutamates of MTX and dihydrofolate inhibit purine and thymidylate biosynthesis


Converted to polyglutamates in normal and malignant tissues. 7-Hydoxylation in liver


t1/2 α = 2 - 3 h; t1/2 β = 8 - 10 hr


Primarily as interact drug in urine

Drug interactions

Toxicity to normal tissues rescued by leucovorin calcium
L-Asparaginase blocks toxicity and antitumor activity
Pretreatment with MTX increases 5-fluorouracil and cytosine arabinoside nucleotide formation
Nonsteroidal anti-inflammatory agents decrease renal clearance and increase toxicity


Mucositis, gastrointestinal epithelial denudation
Renal tubular obstruction and injury


Reduce dose in proportion to creatinine clearance
Do not administer high-dose MTX to patients with abnormal renal function
Monitor plasma concentrations of drug, hydrate patients during high-dose therapy (see Tables 6.2 and 6.4)

t1/2, half-life


In this section, the sequence of events that leads to the cytotoxic action of MTX is considered, beginning with drug movement across the cell membrane, followed by its intracellular metabolism to the polyglutamate derivatives, binding to DHFR and other folate-dependent enzymes, effects on intracellular folates, and, finally, inhibition of DNA synthesis.

Transmembrane Transport

Folate influx into mammalian cells proceeds via two distinct transport systems: (a) the reduced-folate carrier (RFC) system, and (b) the folate receptor (FR) system (Fig. 6.2).31, 32, 33, 34 The proliferative or kinetic state of tumor cells influences the rate of folate and MTX transport. In general, rapidly dividing cells have a greater rate of MTX uptake and a lower rate of drug efflux than cells that are either in the stationary phase or that are slowly growing.35The RFC system, with its large transport capacity, transports folic acid inefficiently (Kt [transport coefficient] = 200 µmol/L) and is a primary transport mechanism of the reduced folates and antifolates like MTX (Kt = 0.7 to 6.0 µmol/L), at pharmacologic drug concentrations.36, 37, 38 The RFC system also transports the naturally occurring reduced folates, including the rescue agent 5-formyltetrahydrofolate (leucovorin).32, 33, 39, 40, 41, 42 Studies have mapped the RFC gene to the long arm of chromosome 21, and this site encodes a protein with predicted molecular size of 58 to 68 kd.43, 44 Several mutations in the RFC have been identified in drug-resistant cell lines, including glutamic acid residue 45, that have been associated with MTX resistance.45, 46, 47 In cells grown under relatively acidic conditions, as may be expected in watershed areas of solid tumor masses, even cell lines with absent RFC have been demonstrated to transport MTX via an RFC-independent pathway48.

Figure 6.2 Transport systems identified for the physiologic folates and various antifolates. (Adapted from Antony AC. The biological chemistry of folate receptors. Blood 1992;79:2807–2820.)

In the second folate transport mechanism, FRs mediate the internalization of folates via a high-affinity membrane-bound 38-kd glycoprotein. The FR gene family encodes three homologous glycoproteins that share a similar folate-binding site. The α and β FRs are anchored to the plasma membrane by a carboxyl-terminal glycosylphosphatidylinositol tail and transport the reduced folates and MTX at a lower capacity than the RFC system. The function of the FR is unknown. The FRs are expressed in normal tissues and, at high levels, on the surface of some epithelial tumors such as ovarian cancer.49, 50 The FR system has a 10-fold to 30-fold higher affinity for folic acid and the reduced folates (Ka [dissociation constant] = 1 to 10 µmol/L) than for MTX. In addition, MTX polyglutamates demonstrate a 75-fold increased affinity for FR compared with the monoglutamate form of MTX.51 Variation in exogenous folate concentrations and normal physiologic conditions, such as pregnancy, can alter the tissue expression of FR. Intracellular levels of homocysteine, which increase under folate-deficient conditions, appear to be the critical modulator of the translational up-regulation of FRs.52 Under conditions of relative folate deficiency, elevated levels of homocysteine stimulate the interaction between heterogeneous nuclear ribinucleoprotein E1 with an 18 base-pair region in the 5′-untranslated region of the FR mRNA resulting in increased translational efficiency, and therefore elevated cellular levels of FR protein. This mechanism serves as an example, in addition to thymidylate synthase and dihydrofolate reductase, of translational efficiency as a mechanism for protein level regulation in the folate biosynthetic pathways.

The FR isoforms (α, β, γ) are independently expressed in mammalian cells and normal human tissues.53 FR-α is expressed in human epithelial neoplasms (ovarian cancer and nasopharyngeal KB carcinoma cells) where it is up- regulated by folate depletion and down-regulated in folate-replete medium.31, 37Elwood et al.54 and other colleagues55, 56 have shown that FR-α is expressed in a complex manner involving promoters upstream from exons 1 and 4 and differential messenger RNA (mRNA) splicing of 5′ exons. FR-β is expressed in human placenta and nonepithelial tumors. FR-γ, found in hematopoietic and lymphatic cells and tissues, lacks a glycosylphosphatidylinositol membrane anchor and is secreted. Although human FR-α, FR-β, and FR-γ share 70% amino acid sequence homology, they differ in binding affinities for stereoisomers of folates.57

Although the precise mechanism of FR-mediated folate uptake remains controversial,58 two separate pathways for FR-mediated folate uptake have been reported: (a) the classic receptor-mediated internalization of the ligand-receptor complex through clathrin-coated pits with subsequent formation of secondary lysosomes, and (b) a mechanism of small molecule uptake, termed potocytosis,59, 60, 61 in which receptor complexes accumulate within distinct subdomains of the plasma membrane known as caveolae that internalize to form intracellular vesicles.62 Once internalization has occurred, acidification within the vesicle causes the folate- receptor complex to dissociate and translocate across the cell membrane. Although questions remain as to the relative importance of the FR and RFC transport systems in the uptake of antifolates during chemotherapy, studies suggest that the RFC system is the more relevant transporter of MTX in mammalian cells, even in cells expressing high levels of FR.63, 64

Both in vitro and in vivo experimental systems have identified defective transport as a common mechanism of intrinsic or acquired resistance to MTX. A number of MTX-resistant cell lines with functional defects in the RFC have now been described.65, 66, 67, 68 An MTX-resistant human lymphoblastic CCRF-CEM/MTX cell line maintained in physiologic concentrations of folate (2 nmol/L) lacked the RFC protein and, for this reason, were resistant to MTX69. These cells retained the folate-binding protein, however, and were able to use this transport process to maintain growth even in nanomolar concentrations of folic acid. This study is of particular interest because the concentration of folate used was in the physiologic range, and thus this mechanism of transport-mediated resistance may have direct clinical relevance.

Zhao et al. have characterized a mutated murine RFC (RFC1) with increased affinity for folic acid and decreased affinity for MTX, which suggests that amino acids in the first predicted transmembrane domain, in particular glutamic acid residue 45, play an important role in determining the spectrum of affinities for, and mobility of, RFC1. This domain is also a cluster region for mutations that occur when cells are placed under selective pressure with antifolates that use RFC1 as the major route of entry into mammalian cells.70, 71 Of interest, however, is that of 121 samples of malignant cells from patients with ALL, none contained a mutation of glutamic acid residue 45.72 To identify clinical MTX resistance on the basis of impaired transport, a sensitive competitive displacement assay using the fluorescent analog of MTX was developed.73 An analysis of 17 patients with acute lymphoblastic leukemia (ALL) revealed that blast cells from two of four patients in relapse after initial treatment with MTX-based combination chemotherapy demonstrated defective MTX transport. In 40 patients with newly diagnosed ALL, low RFC expression at diagnosis was found to correlate with a significantly reduced event-free survival.74 These studies offer evidence that impaired transport may play a role in the development of clinical MTX resistance in patients with ALL. Using semiquantitative reverse-transcription polymerase chain reaction techniques, Guo et al.75 investigated MTX resistance in tumors obtained from patients with high-grade osteosarcoma. In this study, 17 of 26 tumor samples (65%) derived from patients with poor response to chemotherapy had decreased RFC expression. Poor response to MTX-based chemotherapy was also observed in osteosarcoma samples with low levels of RFC at diagnosis.76 These authors concluded that impaired transport of MTX may be a common mechanism of intrinsic resistance in osteosarcoma. An interesting use of the RFC as both a selectable and suicide gene in gene therapy has been recently described.77 Transduction of bone marrow cells with RFC resulted in selection of transfected cells following exposure to trimetrexate, presumably from the increased cellular folate levels and enhanced sensitivity to MTX exposure. Such technology may be useful for enriching cells transduced with various genes that may have therapeutic value. For an in-depth analysis of RFC activity as it relates to transport-mediated MTX resistance, the reader is referred to recent reviews on this subject.78, 79

Significant differences in the characteristics of antifolate drug transport have prompted interest in the development of new analogs. The nonglutamated antifolates such as trimetrexate and piritrexim, as well as the glutamyl esters of MTX, do not require active cellular transport and demonstrate activity against transport-resistant mutants.80, 81 The compound 10-EDAM is more avidly accumulated in tumor cells than in normal bone marrow or intestinal epithelium and has broader therapeutic activity and less marrow toxicity than MTX in experimental systems.25, 82

In contrast to MTX, which has a relatively poor affinity for the folate-binding proteins, several antifolate inhibitors such as CB3717, raltitrexed, DDATHF, pemetrexed (LY231514), and BW1843U89 rely heavily on the high-affinity folate binding proteins for cellular transport.83, 84 Because several of these compounds are efficiently transported by either folate transport system, they may be less susceptible to the emergence of clinical resistance resulting from alterations in membrane transport. Although pemetrexed is transported by both the RFC and FR systems, recent investigations have uncovered an alternative route with a high affinity for this agent in particular.85, 86 This alternative transport system provides a mechanism for continued drug sensitivity even in cells that may have low levels of FR and absent RFC. Homofolate is a DHFR inhibitor that is primarily transported by the folate-binding proteins and has extremely potent activity against malignant cells overexpressing this protein, and thus may be useful for the treatment of human solid tumors that have developed MTX resistance either because of down-regulation or alterations of the RFC system.87 In addition, low-folate conditions that up-regulate FR expression may have clinical relevance. Research has demonstrated that mice maintained on a low-folate diet had higher FR expression on their normal tissues and experienced significantly greater toxicity with the antifolate lometrexol, which suggests the potential for a human corollary in cancer patients with poor nutritional intake.88

Trimetrexate and piritrexim have demonstrated only modest activity against human solid tumors.89, 90 Trimetrexate combined with leucovorin calcium has significant activity against the pulmonary pathogen Pneumocystis carinii, whose DHFR enzyme is highly sensitive to this combination.5, 91 The compound 10-EDAM has clinical activity against a variety of human solid tumors. Its dose-limiting toxicity is mucositis rather than the myelosuppression normally associated with MTX therapy.92 Phase II testing has shown this agent to be active in the treatment of non–small cell lung cancers, soft tissue sarcomas, and breast cancers, with overall response rates of 17%, 14%, and 41%, respectively.93, 94, 95

For many years, at least two poorly defined efflux mechanisms for MTX and the folates have been described including a bromosulfophthalein and probenecid sensitive pathway.96, 97, 98 Recent investigations have identified the multidrug resistance-associated protein family of ATP-binding cassette transporters (MRP-1, -2, -3 and -4) as responsible for cellular MTX efflux with MRP-1 probably representing the primary route.99, 100, 101, 102 Overexpression of this efflux pump has been associated with MTX resistance, and the use of inhibitors such as probenecid have been shown to reverse antifolate resistance.103 Interestingly, it has also been demonstrated that loss of MRP-1 expression may result in antifolate resistance through the expansion of intracellular folate pools, a condition that is well known to be associated with MTX resistance.104 In addition to the MRP family of proteins, the breast cancer resistance protein has also been shown to be associated with the cellular efflux of MTX and MTX polyglutamates with 2 or 3 glutamic acid residues and overexpression of this protein has also been associated with MTX resistance.105, 106, 107

Intracellular Transformation

Naturally occurring folates exist within cells in a polyglutamated form. The polyglutamation of folate substrates is facilitated by folylpolyglutamyl synthetase (FPGS), an enzyme that adds up to four to six glutamyl groups in γ peptide linkage. This reaction serves three main purposes for folates: (a) it facilitates the accumulation of intracellular folates in vast excess of the monoglutamate pool that is freely transportable into and out of cells, (b) it allows selective intracellular retention of these relatively large anionic molecules and thus prolongs intracellular half-life, and (c) it enhances folate cofactor affinity for several folate-dependent enzymes. The MTX polyglutamates are more potent inhibitors of DHFR, TS, AICAR transformylase, and GAR transformylase than is MTX-Glu113, 14 Methotrexate and the other glutamyl-terminal analogs also undergo polyglutamation in normal liver cells, bone marrow myeloid precursors,108, 109human fibroblasts, and a variety of leukemic and carcinoma cell lines.109, 110, 111, 112

The efficiency of the polyglutamation reaction depends on the particular folate substrate and may vary widely among the antifolate compounds. The polyglutamation of MTX occurs over 12 to 24 hours of exposure, at which time most intracellular drug exists in the polyglutamate form.110, 113 In the few studies of polyglutamate formation in vivo, 80% or more of MTX in both normal and malignant tissues was shown to exist in the form of polyglutamates.114,115 Human liver retains MTX polyglutamates for several months after drug administration.116 Thus, the selective retention and depot formation in excess of free monoglutamate, as seen with physiologic folates, appears to characterize MTX polyglutamates as well.

FPGS is a 62-kd magnesium-, adenosine triphosphate–, and potassium-dependent protein.117, 118, 119, 120 The most avid substrate for this enzyme is dihydrofolate (Km [binding affinity] = 2 µmol/L) > tetrahydrofolate (Km = 6 µmol/L) > 10-formyltetrahydrofolate or 5-methyltetrahydrofolate > aminopterin > leucovorin > MTX. Because of the relatively slow rate of formation of MTX polyglutamates compared with the naturally occurring folates, reductions in FPGS activity or cellular glutamate levels that have little effect on folate polyglutamate pools may have critical effects on the level of MTX polyglutamates and on the ultimate cytotoxicity of MTX. Some have postulated that the relatively inefficient metabolism of 5-methyltetrahydrofolate (the predominant folate present in human serum) to its polyglutamate form may be responsible for the folate depletion that occurs in vitamin B12 deficiency. Lack of B12 would inhibit methionine synthetase, which is responsible for the demethylation of 5-methyltetrahydrofolate to tetrahydrofolate, an excellent substrate for FPGS. The accumulation of MTX polyglutamates in liver reduces the polyglutamation of natural folates in that tissue and may, in part, account for the chronic hepatotoxicity associated with MTX. The intracellular content of polyglutamate derivatives represents a balance between the activity of two different enzymes, FPGS and γ-glutamyl hydrolase (GGH, conjugase).121 The latter, a γ-glutamyl-specific peptidase, removes terminal glutamyl groups and returns MTX polyglutamates to their parent monoglutamate form. Models based on leukemic cells from patients with ALL suggest that the terminal one or two glutamic acid residues are most commonly cleaved by hydrolase.122

Yao et al.123 isolated and cloned the complementary DNA (cDNA) for GGH, which codes for an enzyme of 318 amino acids and has a molecular weight of 36 kD. Although it may be expected that overexpression of hydrolase may result in MTX resistance, particularly with brief drug exposures, such was not the case in several human cell line models in which hydrolase was overexpressed.124

MTX polyglutamates exist essentially only within cells and enter or exit cells sparingly.111, 125 The diglutamate form has an uptake velocity of one-fifteenth that of MTX,126 whereas higher glutamates have even slower transport rates.111, 125 Thus, MTX polyglutamates are selectively retained in preference to parent drug as extracellular levels of MTX fall.

Several parameters influence a cell's ability to polyglutamate MTX. Paramount among these factors is the rate of cell growth112, 127 and the level of intracellular folates.127, 128 Enhancement of cell proliferation with growth factors such as insulin, dexamethasone, tocopherol, and estrogen in hormone-responsive cells increases polyglutamation, whereas deprivation of essential amino acids129 results in inhibition of polyglutamation. MTX and L-asparaginase are frequently used in combination for the treatment of acute leukemia. Conversion of MTX to polyglutamate forms can be markedly inhibited by preexposure to L-asparaginase, presumably through amino acid deprivation with resultant growth arrest.130 Increasing intracellular folate pools through exposure of cells to high concentrations of leucovorin or 5-methyltetrahydrofolate results in a decrease in MTX polyglutamation.128 Conversely, the process is enhanced in human hepatoma cells either by incubating cells with MTX in folate-free medium or by first depleting the intracellular folates by “permeabilizing” cell membranes in a folate-free environment.127

An important factor in the selective nature of MTX cytotoxicity may derive from diminished polyglutamate formation in normal tissues relative to that in malignant tissues. Although little metabolism to polyglutamates is observed in normal murine intestinal cells in vivo, most murine leukemias and Ehrlich ascites tumor cells efficiently convert MTX to higher polyglutamate forms in tumor-bearing animals.38, 131 Additionally, normal human and murine myeloid progenitor cells form relatively small amounts of MTX polyglutamates compared with leukemic cells.108, 109

In addition to increasing its retention within cells, polyglutamation of MTX enhances its inhibitory effects on specific folate-dependent enzymes. The pentaglutamates have a slower dissociation rate from DHFR than does MTX132 and a markedly enhanced inhibitory potency for TS (Ki = 50 nmol/L), AICAR transformylase (Ki = 57 nmol/L),163 and, to a lesser extent, GAR transformylase (Ki = 2 µmol/L) in the presence of monoglutamated folate substrates.12, 133The well-described incomplete depletion of physiologic folate cofactors by MTX suggests that direct enzymatic inhibition by MTX polyglutamates may contribute to MTX cytotoxicity. These effects may also explain the competitive nature of leucovorin rescue and the relatively selective rescue of normal versus malignant tissues, in that rescue may depend on the ability of leucovorin and its derived tetrahydrofolates to compete with MTX polyglutamates at sites other than DHFR.

The ability of antifolate analogs to undergo polyglutamation is one of several properties that influences cytotoxic potency. Aminopterin is a better substrate for FPGS than is MTX, and is a more potent cytotoxic agent. A fluorinated MTX analog, PT430, is a weak substrate for FPGS and has little cytotoxic activity.134 The ability to generate polyglutamates has been correlated with sensitivity to MTX and to other antifolate agents that undergo polyglutamation, including pemetrexed and raltatrexed, and is frequently found to be defective in drug-resistant human and murine tumor cell lines,134, 135, 136, 137, 138

Although defective polyglutamation may coexist with other metabolic alterations, examples of pure polyglutamation defects have been described in human leukemia cell lines (CCRF-CEM)139 and in human squamous cancer cell lines derived from head and neck tumors, and have appear to cause of MTX resistance secondary to decreased levels of FPGS.140, 141 Faessel et al.142 evaluated the combined action among polyglutamylatable and nonpolyglutamylatable antifolates directed against various folate-dependent enzymes in human ileocecal HCT-8 cells in vitro and determined that polyglutamation played a critical role in fostering synergy between inhibitors of DHFR and inhibitors of other folate-requiring enzymes. Further evidence for the role of polyglutamation as a determinant of drug sensitivity stems from investigations using other antifolates such as the GAR transformylase inhibitor DDATHF. Polyglutamates of DDATHF were readily formed in cultured human leukemia cell lines and were found to be retained for prolonged periods in drug-free conditions. The FPGS-deficient CCRF-CEM cell line generated few DDATHF polyglutamates and was insensitive to drug exposure.143 Polyglutamation has been investigated as a determinant of response to MTX in clinical chemotherapy. In a study of six human small cell carcinoma cell lines that had demonstrated resistance in vitro after clinical treatment with MTX, two were resistant on the basis of a low capacity to form MTX polyglutamates.144 One of seven samples from MTX-resistant leukemic patients demonstrated a decreased ability to form MTX polyglutamates as the sole explanation for resistance.67 Investigating the reduced accumulation of long-chain MTX polyglutamates in ALL patients, Longo et al.125 found a decrease in the binding affinity (Km) of MTX to FPGS from blast cells of patients with acute myelogenous leukemia (AML) as opposed to ALL. This difference in affinity resulted in a predominance of MTX-Glu1 species in AML cells, and MTX Glu3–5in ALL cells. No corresponding disparity in binding affinity was found when the equivalently cytotoxic antifolate TS inhibitors raltitrexed and BW1843U89, which exhibited similar levels of accumulation of the higher polyglutamate forms, were examined. A more recent study suggests that the evaluation of GGH and folylpolyglutamate synthetase activity at the time of clinical diagnosis may be used as a predictor of the extent of MTX polyglutamation and, therefore, of response to MTX therapy and outcome in patients with acute leukemias.145 Hyperdiploid status in childhood ALL is a good prognostic feature, and patients exhibiting hyperdiploid lymphoblasts show higher levels of synthesis of cytotoxic MTX polyglutamates than patients exhibiting aneuploid or diploid lymphoblasts. Investigators have found a higher concentration of MTX long-chain polyglutamates in T than in B lymphoblasts and an increased level of expression of FPGS mRNA in B-lineage cells.146, 147 These findings suggest that the higher response rates observed in patients with B-cell ALL may result from increased levels of FPGS activity that, in turn, facilitate enhanced intracellular formation of more cytotoxic MTX polyglutamates. However, in a study involving 52 children with B-cell ALL, MTX accumulation and polyglutamation did not appear to have prognostic significance in the context of prolonged oral MTX therapy.148 This study supports the notion that, under the conditions of continuous drug exposure, the activity of MTX may not depend on cellular polyglutamation to sustain intracellular levels. In fresh tumor specimens from patients with soft tissue sarcomas, 12 of 15 patients were determined to be naturally resistant to MTX as a result of impaired polyglutamation.149, 150

Binding to Dihydrofolate Reductase

The physical characteristics of binding of NADPH (reduced form of nicotinamide adenine dinucleotide phosphate [NADP]) and MTX to DHFR have been established by x-ray crystallographic studies, nuclear magnetic resonance spectroscopy, amino acid sequencing of native and chemically modified enzyme, and site-directed mutagenesis. Enzyme from microbial, chicken, and mammalian sources have been studied151, 152, 153, 154, 155, 156; strong amino acid sequence homology is found at positions involved in substrate cofactor and inhibitor binding.157 In general, a long hydrophobic pocket binds MTX and is formed in part by the isoleucine-5, alanine-7, aspartate-27, phenylalanine-31 (Phe-31), phenylalanine-34 (Phe-34), and other amino acid residues. Several particularly important interactions contribute to the binding potency of the 4-amino antifolates: (a) hydrogen bonding of the carbonyl oxygen of isoleucine-5 to the 4-amino group of the inhibitor; (b) a salt bridge between aspartate and the N-1 position of MTX, which is not involved in binding to the physiologic substrates; (c) hydrophobic interactions of the inhibitor with DHFR, particularly with Phe-31 and Phe-34; (d) hydrogen bonding of the 2-amino group to aspartate-27 and to a structurally consistent bound water molecule; and (e) hydrogen binding of the terminal glutamate to an invariant arginine-70 residue. Investigations have identified the importance of the interactions of MTX with Phe-31 and Phe-34 because mutations in these positions result in a 100-fold and 80,000-fold decrease in MTX affinity for the enzyme, respectively.158 Mutation of arginine-70 results in a decrease in MTX affinity by >22,000-fold but does not alter the binding affinity of trimetrexate, which lacks the terminal glutamate moiety. This finding supports the essential role of arginine-70 in the binding of inhibitors that preserve the terminal glutamate structure.159 Mutations outside the enzyme active site also may result in marked reductions in folate and antifolate affinities.160 In addition, the physiologic substrate dihydrofolate is bound to the enzyme in an inverted, or “upside down,” configuration compared with the inhibitor MTX.155, 161 The reader is referred to more detailed reviews of this subject for consideration of substrate and cofactor binding characteristics and mutated DHFR cDNA studies.153, 154, 155, 156, 162, 163, 164, 165

Optimal binding of MTX to DHFR depends on the concentration of NADPH. NADH (reduced form of nicotinamide adenine dinucleotide) may also act as a cosubstrate for DHFR but, unlike NADPH, it does not promote binding of MTX to the enzyme.166 Thus, the intracellular ratios of NADPH/NADP and NADPH/ NADH may play an important role in the selective action of MTX to the extent that the cosubstrate ratios may differ in malignant and in normal tissues.131, 166In the presence of excess NADPH, the binding affinity of MTX for DHFR has been estimated to lie between 10 and 200 pmol/L,167, 168 although this affinity is significantly affected by pH, salt concentration, and the status of enzyme sulfhydryl groups. Under conditions of low pH and with a low ratio of inhibitor to enzyme, binding is essentially stoichiometric, that is, one molecule of MTX is bound to one molecule of DHFR.

Binding of MTX to DHFR isolated from bacterial and mammalian sources in the presence of NADPH generates a slowly formed ternary complex. The overall process has been termed slow, tight-binding inhibition and involves an initial rapid but weak enzyme-inhibitor interaction followed by a slow but extremely tight-binding isomerization to the final complex.153, 165, 169 The final isomerization step probably involves a conformational change of the enzyme with subsequent binding of the para-aminobenzoyl moiety to the enzyme.154 Other folate analogs, such as aminopterin, follow the same slow, tight-binding kinetic process, in contrast to the pteridines and pyrimethamine, which behave as classic inhibitors of the bacterial enzymes. Trimethoprim is considered to be a classic, albeit weak, inhibitor of mammalian DHFR. Of note, it does not undergo an isomerization process to the ternary complex form.165

In the therapeutic setting, MTX acts as a tight-binding but reversible inhibitor. Under conditions of high concentrations of competitive substrate (dihydrofolate) and at neutral intracellular pH, a considerable excess of free drug is required to fully inhibit the enzyme. Both in tissue culture and in cell-free systems, tritium-labeled MTX bound to intracellular enzyme can be displaced by exposure of cells to unlabeled drug, dihydrofolate,7, 170, 171 or reduced folates such as leucovorin and 5-methyltetrahydrofolate,131 which indicates a slow but definite “off rate” or dissociation of MTX from the enzyme.131, 172 Thus, an excess of free, or unbound, drug is required to maintain total inhibition of DHFR.173

The polyglutamates of MTX have similar potency in their tight-binding inhibition of mammalian DHFR110, 165, 174 and possess a slower rate of dissociation from the enzyme than the parent compound. In pulse-chase experiments using intact human breast cancer cells, MTX pentaglutamate was found to have a dissociation half-life of 120 minutes compared with 12 minutes for the parent compound. Cell-free experiments using purified preparations of mammalian enzyme indicate that MTX polyglutamation has a modest effect in enhancing binding and catalytic inhibition (twofold to sixfold) of DHFR.12, 165, 168, 175 As with MTX, enzyme-bound MTX polyglutamates may also be displaced by reduced folates131 and high concentrations of dihydrofolate,176, 177 albeit at a slower rate than MTX.

These observations indicate that, in the absence of free drug, a small fraction of intracellular DHFR, either through new synthesis or through dissociation from the inhibitor, becomes available for catalytic activity and is adequate to allow for continued intracellular metabolism. The requirement for excess free drug to inhibit enzyme activity completely is important in understanding the clinical effects and toxicity of this agent, and is fundamental to the relationship between pharmacokinetics and pharmacodynamics.

Resistance to MTX as a result of decreased DHFR binding affinity for MTX has been described in murine leukemic cells,160, 178, 179 Chinese hamster ovary180and lung181 cells, and murine and hamster lung fibroblast cells.182, 183 These mutant enzymes may have several thousand–fold reduced binding affinity for MTX and, in general, are less efficient in catalyzing the reduction of dihydrofolate than is wild-type DHFR.

Drug-sensitive Chinese hamster lung cells have been found to express two different forms of DHFR encoded by distinct alleles.184, 185 The two species differ in molecular weight and isoelectric point (21,000 versus 20,000 and 6.7 versus 6.5) and result from a single amino acid substitution of asparagine for aspartic acid at position 95. Either allele may be predominantly expressed in various subclones of the parent cell line. This observation raises the possibility that distinct naturally occurring DHFR alleles may exist in a variety of tissues and, to the extent that they may confer differential sensitivity to MTX, this DHFR genetic polymorphism of the host may serve as a mechanism by which cells may become clinically MTX resistant.

Figure 6.3 A. Marker chromosomes found in methotrexate (MTX)-resistant breast cancer cells. A human breast cancer cell line, MCF-7, resistant to MTX was isolated by growing cells in gradually increasing drug concentrations. These cells are resistant to drug concentrations more than 200-fold higher than those that kill wild-type cells and contain more than 30-fold increases in dihydrofolate reductase (DHFR). The arrow indicates a marker chromosome with a greatly expanded homogenously staining region. (Courtesy of National Cancer Institute, Bethesda, MD) B. Metaphase plate of a small cell lung cancer carcinoma cell line taken from a patient with clinical MTX resistance. The prominent double-minute chromosomes (arrows) were associated with amplification of the drug target enzyme, DHFR. (From Curt GA, Carney DN, Cowan KH, et al. Unstable methotrexate resistance in human small-cell carcinoma associated with double minute chromosomes. N Engl J Med 1983;308:199–202.)

DHFR with reduced affinity for MTX may represent a clinically important mechanism of MTX resistance, as this phenomenon was observed in the leukemic cells of 4 of 12 patients with resistant AML.186 MTX-resistant mutant DHFR has been used as a means to protect and/or select transduced hematopoietic progenitor cells and several laboratories have developed vectors to efficiently transduce human progenitor cells with the hope that such technology could be used to enable the use of higher, and hopefully more effective, doses of chemotherapeutic agents to treat human maligancies.187, 188 A common finding in MTX-resistant cells is an increase in the expression of DHFR protein with no associated change in the enzyme's affinity for MTX. Elevations in DHFR may persist for many generations of cell renewal in tumor cells from resistant patients. In resistant murine leukemic cells, the increased DHFR activity results from reduplication of the DHFR gene (Fig. 6.3), a process that has been shown to occur by exposing murine and human leukemia and carcinoma cells in culture to stepwise increases in the concentration of MTX.181, 184, 189, 190, 191 Gene reduplication may take the form of a homogeneously staining region (HSR) on chromosomes or nonintegrated pieces of DNA known as double-minute chromosomes (Fig. 6.3B). Although HSRs appear to confer stable resistance to the cell, double-minute chromosomes are unequally distributed during cell division,183, 190 and in the absence of the continued selective pressure of drug exposure, the cells revert to the original low-DHFR genotype. Evidence exists that gene amplification occurs initially in the form of double-minute chromosomes because this is the predominant abnormality in low-level drug-resistant cells, whereas HSRs occur in highly resistant cells that contain multiple gene copies.183, 190, 191 Other investigations suggest the opposite sequence wherein chromosomal breaks result in HSRs, which are then processed to DMs or not, depending on how different cell types handle extra chromosomal sequences.192 Another mechanism of gene amplification has been identified in an MTX-resistant HeLA 10B3 cell line in which were found submicroscopic extrachromosomal elements (amplisomes) containing amplified DHFR genes. These amplisomes appeared early in the development of MTX resistance and were not found to be integrated into the chromosome, nor were they associated with double- minute chromosomes. Although these amplisomes were lost in the absence of the selective pressure of MTX, they disappeared at a much slower rate than would be predicted from simple dilution of nonreplicating elements.

Although MTX resistance through DHFR gene amplification becomes apparent only after the prolonged selective pressure of drug exposure, studies indicate that highly MTX-resistant cells may be generated by gene amplification within a single cell cycle.193 Early S-phase cells exposed transiently to agents that block DNA synthesis (e.g., hydroxyurea) may undergo reduplication of multiple genes synthesized during early S phase, including DHFR, after removal of the DNA synthetic inhibitor. This finding has broad implications for the rapid development of drug resistance in patients treated with MTX and other inhibitors of DNA synthesis. Exposure of cells to a variety of chemical and physical agents unrelated to MTX including hypoxia, alkylating agents,194 ultraviolet irradiation,194, 195 phorbol esters,194, 195, 196 cis-diamminedichloro-platinum,197 doxorubicin,198 and 5-fluorodeoxyuridine199 may induce MTX resistance through DHFR gene amplification, with subsequent increases in DHFR protein. The induction of MTX resistance by a variety of chemical and physical agents may explain de novo MTX resistance in certain human tumors, given the constant presence of a host of environmental carcinogens. Unlike malignant cells, amplification of DNA has not been reported in normal cells of patients undergoing therapy with cytotoxic agents or in cell lines of normal cells.200

In addition to gene amplification, more subtle mechanisms exist for increasing DHFR expression. Molecular analysis of the DHFR gene encoding for overexpressed DHFR protein has occasionally revealed significant differences in non–protein coding regions that may impact mRNA expression.184 The E2F-1 transcription factor has been shown to promote the transcription of DHFR as well as thymidylate synthase mRNA and has been correlated with the messenger RNA levels of these two enzymes in tumor samples of patients with osteosarcoma.201 Exposure of human breast cancer cells to MTX results in an acute increase (up to fourfold) in the cellular DHFR content.202 The expression of DHFR protein in this setting appears to be controlled at the level of mRNA translation, as no acute associated change occurs in the amount of DHFR mRNA or DHFR gene copy number after MTX exposure nor are alterations seen in DHFR enzyme stability. Using an RNA gel mobility shift assay, human recombinant DHFR protein was shown to specifically bind to its corresponding DHFR mRNA.203 Incubation of DHFR protein either with the normal substrates dihydrofolate or NADPH, or with MTX, completely represses its binding to the target DHFR mRNA. In an in vitro translation system, this specific interaction between DHFR and its message is associated with inhibition of translation. These studies provide evidence for a translational autoregulatory mechanism underlying the control of DHFR expression. The presence of either excess MTX or dihydrofolate prevents DHFR protein from performing its normal autoregulatory function, thereby allowing for increased DHFR protein synthesis. Thus, the ability to regulate DHFR expression at the translational level allows normal cellular function to be maintained in the setting of an acute cellular stress and represents a unique mechanism whereby cells can react to and overcome the inhibitory effects of MTX and antifolate analogs. In further attempts to characterize DHFR autoregulation, Bertino et al.204 used a series of truncated DHFR mRNA probes to investigate whether the enzyme directly contacts its cognate mRNA. A resultant DHFR protein/RNA interaction was found in an approximately 100–base-pair portion in the protein-coding region that contains two putative stem-loop structures. In addition, the binding of MTX prevented the DHFR/RNA interaction to DHFR, which thereby relieved translational autoregulation.

Although various in vitro and in vivo model systems have clearly demonstrated an association between DHFR gene amplification and MTX resistance, the clinical significance of gene amplification remains uncertain. Tumor samples from patients resistant to MTX have been evaluated, and several clinical specimens have been found to possess elevated levels of DHFR enzyme in association with DHFR gene amplification.205 A small cell lung carcinoma cell line isolated from a patient clinically resistant to high-dose MTX was found to have amplification of the DHFR gene and increased expression of DHFR protein.205,206 This amplification was associated with the presence of double-minute chromosomes (Fig. 6.3B). After serial passage in drug-free media, cells lost the double-minute chromosomes on which the amplified genes resided and regained drug sensitivity. Clinical MTX resistance attributable to DHFR amplification was also investigated in two patients with acute leukemia and in one patient with ovarian cancer. In all three cases, amplification of DHFR gene copies (twofold to threefold) with increased DHFR protein (threefold to sixfold) was observed, and the increase in DHFR gene copy number was postulated to be directly associated with the development of MTX resistance. Matherly et al.207 found a markedly greater frequency of DHFR overexpression in T-cell ALL than in B-precursor ALL in children. The authors speculated that this difference in DHFR expression was associated with the poorer prognosis of T-cell ALL treated with standard doses of antimetabolites, implying that higher-dose MTX consolidation therapy may be particularly needed in this population.

Consequences of Dihydrofolate Reductase Enzyme Inhibition

The critical cellular events associated with MTX inhibition of DHFR are illustrated in Figure 6.1. Thymidylate synthase catalyzes the sole biochemical reaction resulting in the oxidation of tetrahydrofolates. Continued activity of this enzymatic reaction in the presence of DHFR inhibition results in rapid accumulation of intracellular levels of dihydrofolate polyglutamates. This accumulation is temporally associated with a depletion of several critical reduced-folate pools, most notably 5-methyltetrahydrofolate. In several in vitro cell systems studied to date, however, the reduced-folate cofactors required for de novo purine and thymidylate synthesis (10-formyltetrahydrofolate and 5,10-methylenetetrahydrofolate) are relatively preserved in the presence of cytotoxic concentrations of MTX.7, 8, 9, 10, 12, 208 In studies using human breast cancer cells, purified normal human myeloid precursor cells, and murine leukemia cells, exposure to lethal concentrations of MTX resulted in a 70 to 80% preservation of 10-formyltetrahydrofolate pools compared with untreated controls (Fig. 6.4). Additional studies using human breast cancer cells, promyelocytic leukemia cells, normal human myeloid progenitor cells, and Krebs ascites cells and L1210 murine leukemia cells grown in the peritoneal cavity of mice confirm a partial preservation of 5,10- methylenetetrahydrofolate pools (50 to 70%) during MTX exposures that produce profound TS inhibition and cytotoxicity.11, 12 Subsequent computer modeling of the intracellular human folate pools based on experimental data confirm the importance of direct inhibition of the various folate-dependent enzymes in the metabolic inhibition associated with MTX exposure.20 To determine the importance of the inhibitory effects of MTX polyglutamates as distinct from the effects of folate depletion and direct dihydrofolate inhibition, several investigators have examined the change in folate pools associated with exposure to trimetrexate, an antifolate that is not polyglutamated but remains a potent inhibitor of DHFR.20, 209, 210, 211 Results of these studies are conflicting. In murine leukemia cells, folate pools were relatively preserved and dihydrofolate polyglutamates seemed to serve an essential role in metabolic inhibition. In contrast, folate depletion appeared to be the more critical event in rat hepatoma cells. Presumably, intrinsic differences between these cell lines with regard to the levels of TS, folate pools, and intracellular folate regulation may determine the relative roles of direct enzyme inhibition versus substrate depletion in the metabolic inhibitory effects of MTX. Although partial depletion of reduced-folate cofactors undoubtedly contributes to the inhibition of metabolic pathways, the accumulated dihydrofolate and MTX polyglutamates appear also to play a critical role as direct inhibitors of folate-dependent enzymes in both de novo purine and thymidylate synthesis.

Figure 6.4 Effects of 1 µmol/L methotrexate (MTX) on intracellular folate pools in human breast cancer cells (MCF-7). (Δ, dihydrofolate; ○, 10-formyldihydrofolate; ▪, 10-formyltetrahydrofolate; ◊, 5-methyltetrahydrofolate.) (From Allegra CJ, Fine RL, Drake JC, et al. The effect of methotrexate on intracellular folate pools in human MCF-7 breast cancer cells. Evidence for direct inhibition of purine synthesis. J Biol Chem 1986;261:6478–6485.)

Reduced-folate (leucovorin) rescue of MTX-treated cells may be anticipated to result in an accumulation of reduced folates that would compete with and overcome direct enzymatic inhibition rather than simply replete tetrahydrofolate levels (vide infra). This feature may, in large part, explain the competitive nature of leucovorin rescue observed in vitro and clinically. Also, selectivity of the cytotoxic effects of MTX and selectivity of leucovorin rescue may depend on the extent to which various normal and malignant cells generate dihydrofolate and MTX polyglutamates.

An additional factor that influences the folate pool changes associated with MTX exposure and, hence, cellular sensitivity to MTX is the level or activity of TS. Inhibition of TS by 5-fluorodeoxyuridylate or by depletion of its substrate deoxyuridylate diminishes sensitivity to MTX. TS activity in human leukemia, lung carcinoma, and colon carcinoma influences MTX sensitivity; specifically, low levels of TS activity are usually associated with MTX resistance.67, 144 In cells with low levels of TS, the slow rate of oxidation of 5,10-methylenetetrahydrofolate to dihydrofolate creates less dependence on DHFR to regenerate tetrahydrofolates. Under conditions of low cellular TS activity, a block in DHFR by MTX exposures produces minimal accumulation of inhibitory dihydrofolate and minimal depletion of tetrahydrofolates.

Mechanisms of Cell Death

As a consequence of the multiple effects of antifolates on nucleotide biosynthesis, several mechanisms of cell death are possible. Deoxythymidine triphosphate and deoxypurine nucleotides are required for both the synthesis of DNA and its repair. Inhibition of thymidylate and purine synthesis leads to a cessation of DNA synthesis. A close correlation is found between DNA strand breaks and cell death in Ehrlich ascites tumor cell exposed to MTX212; because the breaks occurred in mature DNA, the authors attributed them to ineffective repair mechanisms from lack of nucleotides. This work was supported by similar experimental findings in a mutant murine cell line lacking TS activity and grown in thymidine-deplete media213 and in a Chinese hamster ovary cell line in which DNA damage was prevented by the use of thymidine and hypoxanthine.214 Another hypothesis that attempts to explain MTX cytotoxicity concerns the increase in intracellular dUMP pools that occurs as a consequence of inhibition of de novo thymidylate synthesis. Clearly, the high concentrations of dUMP may ultimately lead to misincorporation of dUMP and deoxyuridine triphosphate into cellular DNA.215 An enzyme, uracil-DNA-glycosylase, specifically excises uracil bases from DNA, a process that may be responsible for the fragments of DNA observed in antifolate-treated cells with high levels of uracil incorporation.216,217 These studies implicate the presence of lesions expected in DNA undergoing excision repair of misincorporated uracil nucleotides. Further evidence for the importance of uracil misincorporation derives from a study of seven cell lines that varied widely in deoxyuridine triphosphatase activity.218 An inverse correlation between deoxyuridine triphosphatase activity and MTX toxicity was found, which suggests that the level of deoxyuridine triphosphate misincorporation into DNA is an important factor in MTX cytotoxicity. Although these studies offer insights into the consequences of uracil misincorporation into DNA, they do not explain the marked toxicity of MTX-thymidine combinations, which must act through an antipurine effect. Probably both uracil nucleotide misincorporation (with subsequent excision repair) and the combined effects of purine and pyrimidine depletion result in the formation of DNA strand breaks. Although the induction of DNA strand breaks is central to the activity of MTX, it is the cellular response to these breaks that ultimately determines whether a cell incurring a given level of DNA damage dies.219, 220, 221

Using a tetracycline-inducible expression system in an osteosarcoma cell line (SaOs-2), Li et al.222 found that p21/waf1-induced cells exhibited greater sensitivity to doxorubicin hydrochloride, raltitrexed, and MTX than noninduced cells and that this condition was associated with increased apoptosis. The SaOs-2 cells lack both p53 and a functional retinoblastoma protein. Overexpression of p21/waf1 protein was associated with diminished E2F-1 phosphorylation, which resulted in an increase in E2F-1 binding activity and enhanced expression of E2F-responsive genes (DHFR and TS). The authors suggested that this mechanism may mediate sensitivity to anticancer drugs by contributing to increased S–G2 cell-cycle arrest or delay and increased cell susceptibility to apoptosis. Clearly, the identification of the critical cell death effectors will enable the development of more potent and, it is hoped, more selective therapeutic strategies.

A novel antiproliferative mechanism associated with MTX exposure has been described as resulting from the potent inhibition of isoprenylcysteine carboxyl methyltransferase (Icmt).223 Inhibition of Imct results from the elevated levels of S-adenosylhomocysteine that occur with MTX exposure. Icmt is responsible for the Ras methylation, which is necessary for its proper function and, inhibition of this methylation process results in proliferative arrest. The relative extent to which this process contributes to the antiproliferative effects of MTX is not clear.

Pharmacokinetic and Cytokinetic Determinants of Cytotoxicity

At least two pharmacokinetic factors—drug concentration and duration of cell exposure—are critical determinants of cytotoxicity. In tissue culture and in intact animals, extracellular drug concentrations of 10 nmol/L are required to inhibit thymidylate synthesis in normal bone marrow. This same drug concentration is associated with depletion of bone marrow cellularity when maintained for 24 hours or longer. The rate of cell loss from murine bone marrow increases with increasing drug concentrations up to 10 µmol/L224 (Fig. 6.5). Similar findings have been reported in studies with murine tumor cells. Compared with drug concentration, the duration of exposure to MTX is a more critical factor in determining cell death, provided the minimal threshold concentration for cytotoxicity is exceeded. For a given dose of drug, cell loss is directly proportional to the time period of exposure but doubles only with a 10-fold increase in drug concentration.225 This relationship is likely the result of the S-phase specificity of MTX. With longer duration of exposure, more cells are allowed to enter the vulnerable DNA-synthetic phase of the cell cycle.

Time and concentration correlates of cytotoxicity for human tumor cells have also been studied. Hryniuk and Bertino226 found variable inhibition of thymidine incorporation into DNA of human leukemic cells in short-term culture when these cells were exposed to 1 µmol/L MTX for 1 hour or less. In leukemia cell lines, the duration of exposure appears to play a far more important role than absolute drug concentration. A marked increase in toxicity (30-fold) was appreciated only when the duration rather than drug concentration was increased,225 provided both parameters were changed by similar increments.

Figure 6.5 Nucleated cells per femur remaining after constant infusion of methotrexate sodium into mice to achieve indicated drug concentration for various periods. (From Pinedo HM, Zaharko DS, Bull J, et al. The relative contribution of drug concentration and duration of exposure to mouse bone marrow toxicity during continuous methotrexate infusion. Cancer Res 1977;37:445–450.)


In addition to pharmacokinetic and cytokinetic factors, physiologic compounds in the cellular environment may profoundly affect the cytotoxicity of MTX. Most prominent among these factors are the naturally occurring purine bases, purine nucleosides, and thymidine. In bone marrow and intestinal epithelium, the de novo synthesis of both thymidylate and purines is inhibited by concentrations of MTX above 100 nmol/L, but cells can survive this block when bone marrow is supplied with 10 µmol/L thymidine and a purine source (adenosine, inosine, or hypoxanthine) at similar concentrations. Thymidine alone is incapable of completely reversing the cytotoxic effect of MTX.227 The purine salvage pathways in normal bone marrow appear to be highly efficient, however, and the endogenous concentrations of purines in this tissue are high, albeit variable.228 Plasma thymidine levels in humans have been reported to be approximately 0.2 µmol/L,229 whereas the concentration of the purine bases and nucleosides is somewhat higher (0.5 µmol/L).230 Thus, under basal conditions, the concentrations of purines and thymidine would appear inadequate to rescue cells. Clinical investigations using the nucleoside transport inhibitor dipyridamole, however, have demonstrated an increase in toxicity when combined with MTX, which suggests that physiologic concentrations of nucleosides may affect the toxicity and potentially the antitumor activity of MTX.231 Recent work has demonstrated that physiologic levels of endogenous nucleosides and nucleobases are adequate to sensitize even highly resistant osteosarcoma cell lines to the cytotoxic effects of MTX.232 Pharmacologic interventions, such as allopurinol treatment (which elevates circulating hypoxanthine concentrations) and chemotherapy, with subsequent tumor lysis, may further raise levels of the circulating nucleosides and ameliorate toxicity to tumor or host tissues.

A third determinant of antifolate cytotoxic action is the concentration of reduced folate in the circulation. Methyltetrahydrofolate (the predominant circulating folate cofactor), when present in sufficient concentration, can readily reverse MTX toxicity,233 as can leucovorin. Circulating levels of 5-methyltetrahydrofolate are approximately 0.01 µmol/L and of little pharmacologic relevance. Exogenous administration of reduced folates, however, is able to reverse MTX toxicity in a competitive manner. Leucovorin is commonly used after MTX administration to reduce or prevent toxicity and is effective when given within 24 to 36 hours after MTX treatment. The concentration of leucovorin required to prevent MTX toxicity increases as the drug concentration increases (Fig. 6.6).227 The reasons for this competitive relationship are only partly understood. However, given the current knowledge of direct enzymatic inhibition of the folate-dependent enzymes in de novo purine and thymidylate synthesis by intracellular metabolites formed after MTX treatment, several possibilities exist. Competition may occur at the level of membrane transport because both MTX and the reduced folates share a common transmembrane transport system. When given concurrently with MTX, leucovorin decreases the rate of MTX polyglutamation. An important consideration in leucovorin rescue is its effect on intracellular reduced-folate pools. Competitive concentrations of reduced folate are required to overcome inhibition by dihydrofolate and MTX polyglutamates as folate- dependent enzymes such as TS and AICAR transformylase. In addition, by raising intracellular concentrations of dihydrofolate, leucovorin indirectly increases substrate competition with MTX for the inhibited DHFR enzyme. In support of this latter mechanism as the basis for leucovorin rescue are studies that show nearly quantitative conversion of leucovorin to dihydrofolate and the ability of dihydrofolate to (a) compete with MTX and MTX polyglutamates for DHFR binding in intact human cancer cells, (b) reactivate DHFR, and (c) rescue cells from MTX cytotoxicity.7, 171, 177, 234 Furthermore, an important factor in the clinical efficacy of leucovorin rescue relates to the timing and dose of leucovorin.235 Because leucovorin is capable of reversing the cytotoxic effects of MTX on host and malignant cells, the minimum dose of leucovorin needed to rescue host cells should be used. In patients with head and neck cancer randomized to receive standard-dose MTX and either leucovorin or placebo rescue starting 24 hours later,236 both overall toxicity and response rate were significantly lower in patients treated with MTX and leucovorin. This study emphasizes that careful consideration must be given to the dose and timing of leucovorin when used in combination with MTX to avoid rescue of both cancerous and normal tissues.

Figure 6.6 Effect of various combinations of leucovorin calcium and methotrexate sodium (MTX) on formation of granulocyte colonies in vitro by mouse bone marrow. Values are normalized to control value for marrow incubated without either drug. (MTX concentrations: ▪, 10-9 mol/L; □, 10-8 mol/L; Δ, 10-7mol/L; ○, 10-6 mol/L; ▲, 10-5 mol/L; ●, 10-4 mol/L.) (From Pinedo HM, Zaharko DS, Bull JM, et al. The reversal of methotrexate cytotoxicity to mouse bone marrow cells by leucovorin and nucleosides. Cancer Res 1976;36:4418–4424.)



Two methods are commonly used for the rapid assay of MTX and include one that is based on tight binding of drug to DHFR and another that depends on antibody-drug interactions. Both methods provide extremely sensitive measurement of MTX levels >10 nmol/L in biologic fluids. Significant differences are seen, however, in the time required to perform them and in their specificity for parent compound as opposed to metabolites (Table 6.2).

The first widely used method, the enzyme inhibition assay, measures MTX concentration by determining the ability of a clinical sample to inhibit DHFR activity. Although the enzyme inhibition assay is time-consuming, automation and microcomputer technology may allow for a more efficient use of this very sensitive and specific assay.237

A competitive binding assay that uses DHFR as the binding protein is available and preserves the specificity for parent compound that is observed in the enzyme inhibition assay. In the binding assay, MTX in a biologic sample competes for DHFR binding sites with a known quantity of radiolabeled drug. Multiple samples can be run simultaneously and results can be reported on the same day.238 Trimethoprim, which is commonly used in the oncologic population in combination with sulfonamides for the treatment and prophylaxis of P. carinii pneumonia and bacterial infections, cross-reacts with MTX in this assay and may lead to spuriously elevated results when the bacterial enzyme is used.239 This cross-reaction may be avoided by using a mammalian source of DHFR (e.g., bovine) that does not bind trimethoprim at clinically achievable serum concentrations.





DHFR inhibition

Sensitive, no cross-reaction with metabolites


Competitive DHFR binding

Sensitive, no cross-reaction with metabolites, rapid

Not automated

   Fluorescence polarization

Sensitive, rapid, automated

Cross-reacts with metabolites

   Enzyme multiplied

Rapid, automated

Relatively insensitive, cross-reacts with metabolites

   High-pressure liquid chromatography

Methotrexate and metabolites individually quantitated

Time consuming, expensive

DHFR, dihydrofolate reductase.

Radioimmunoassays and fluorescence-polarization immunoassays for MTX are also available for routine clinical use, and employ antibodies generated by an MTX-bovine serum albumin complex.240 These assays are as sensitive (0.01 to 800 µmol/L) and as rapid as the competitive DHFR binding procedure but have somewhat different specificity.
The immunoassay antibodies cross-react with one MTX metabolite, 2,4-diamino-N10-methyl pteroic acid (DAMPA) (40%), but not with 7-hydroxymethotrexateMTX (7-OH-MTX) (1%). At later time points after drug administration, DAMPA is found in plasma in relatively high concentrations (equal to or greater than that of MTX) and thus produces spuriously elevated values for the parent compound241 that are on average twofold to fourfold higher than the DHFR binding method. A variant of the radioimmunoassay (enzyme-multiplied immunoassay), based on antibody inhibition of an enzyme-MTX complex, is the most rapid assay available but also cross-reacts with DAMPA242 and has a sensitivity limit of approximately 0.2 µmol/L. This limitation is critical when evaluating patient samples beyond 48 hours, when MTX concentrations may be at the limit of the assay's sensitivity. Prudent use of hydration and continued leucovorin can be guided only by accurate measurements of MTX concentrations down to the nontoxic level (approximately 10 nmol/L).243

High-pressure liquid chromatography (HPLC) can be used to separate and quantitate MTX and its various metabolites.241, 244 Although the sensitivity of HPLC is primarily limited by the ultraviolet detection systems commonly used (0.2 µmol/L), its sensitivity may be markedly enhanced by serum concentration methods,244, 245 electrochemical detection,246 or post column derivatization and fluorescence detection. Although HPLC technology is generally too cumbersome for routine clinical monitoring, its use is required when high sensitivity and specificity along with an ability to measure individual MTX metabolites are required. Reports have compared the two more commonly used assay techniques, enzyme-multiplied immunoassay and fluorescence-polarization immunoassay, with HPLC and in over 100 patient plasma samples tested found a high concordance among all three methods.247 Thus, several assays are available for the accurate and rapid measurement of MTX levels in patient samples. The final selection of an assay to be used in the clinical setting may ultimately depend on the requirements for sensitivity, specificity, identification of specific MTX metabolites, and cost and time constraints.


Although the pharmacologic properties of MTX are now better understood at the biochemical and molecular levels, clinical exploitation of this knowledge depends on a detailed understanding of the time profile of drug concentration in extracellular and intracellular spaces and the complex relationship of drug levels to effects on specific tissues. Although antifolate pharmacokinetics is well understood, the important second step—that is, definition of the relationship between drug concentration and effect (pharmacodynamics)—requires continued investigation.

The first attempts to define the distribution and disposition of MTX in a comprehensive manner were reported by Zaharko et al.248 These authors developed a detailed model for MTX pharmacokinetics that accurately predicted drug-derived radioactivity in various tissue compartments for a 4-hour period after drug administration. The primary elements of that model were (a) elimination of MTX by renal excretion, (b) an active enterohepatic circulation, (c) metabolism of at least a small fraction of drug within the gastrointestinal tract by intestinal flora, and (d) multiple drug half-lives in plasma, the longest of which was found to be approximately 3 hours. Each of these elements has been observed in humans, although a longer terminal half-life is now appreciated, estimated to be between 8 and 27 hours, depending on the assay method used.248 HPLC has also disclosed extensive metabolism of MTX in both mice and humans.


MTX is absorbed from the gastrointestinal tract by a saturable active transport system.249 Although small doses are well absorbed, absorption is incomplete at higher doses. Bioavailability for doses of 50 mg/m2 or greater may be enhanced by subdividing the dose rather than delivering as a single large dose.249, 250An investigation of the oral bioavailability of MTX given to 15 patients with acute leukemia (6 to 28 mg/m2) revealed a longer absorptive phase and a lower fractional absorption of the drug for doses >12 mg/m2 (2.5 hours and 51%, respectively) than for doses less than 12 mg/m2 (1.5 hours and 87%, respectively).251

Drug absorbed in the intestine enters the portal circulation and thus must pass through the liver, where hepatocellular uptake, polyglutamation, and storage all occur; orally administered drug is further subject to degradation (deglutamylation) by intestinal flora to DAMPA, a metabolite that is inactive pharmacologically but that cross-reacts in commonly used radioimmunoassay systems for MTX. Drugs taken orally are also subject to the variability of intestinal absorption created by drug-induced epithelial damage, motility changes, and alterations in flora. One or all of these factors may be responsible for the highly variable nature of MTX pharmacokinetics observed in children receiving small doses of MTX (20 mg/m2) by mouth.252 For these reasons, MTX is usually given by systemic routes.


The volume of distribution of MTX approximates that of total body water. The drug is loosely bound to serum albumin, with approximately 60% binding at concentrations at or above 1 µmol/L in plasma.253 Weak organic acids such as aspirin254 can displace MTX from plasma proteins, but the clinical significance of this displacement process has not been proven.

MTX penetrates slowly into third-space fluid collection, such as pleural effusions255 or ascites,256 reaching steady-state plasma concentrations in approximately 6 hours. It also exits slowly from these compartments, producing a concentration gradient of several-fold in favor of the loculated fluid at later time points. The clearance of MTX from peritoneal fluid is approximately 5 mL/minute, substantially less than its clearance from the plasma compartment, which equals or exceeds glomerular filtration (120 mL/minute). In brief, the mechanism responsible for drug accumulation in closed fluid spaces relates to the limited permeability of the peritoneal surface to both charged and high-molecular-weight compounds257; thus, only small amounts of drug are able to cross the peritoneal membrane and enter the portal circulation. Drug either not retained or metabolized in the liver passes into the systemic circulation and is then excreted rapidly in the urine.

Third-space retention of intravenously administered drug is associated with a prolongation of the terminal drug half-life in plasma, owing presumably to the slow reentry of sequestered drug into the bloodstream.256 This effect must be considered when treating patients with ascites or pleural effusions. Although no strict guidelines exist for dose adjustment in patients with third-space accumulations, evacuation of this fluid before treatment and close monitoring of plasma drug concentrations in such patients is strongly advisable.

Fossa et al.258 have described unexpectedly high levels of MTX in patients with bladder cancer receiving MTX-based combination chemotherapy who had previously undergone cystectomy and ileal conduit diversion, presumably because of drug resorption from the ileal conduit. Thus, special care must be given to patients with ileal conduit diversions who are at increased risk for delayed MTX elimination and subsequent MTX toxicity.

Plasma Pharmacokinetics

After the initial distribution phase, which lasts a relatively few minutes, at least two phases of drug disappearance from plasma are observed in laboratory animals and humans. Conventional doses of 25 to 100 mg/m2 produce peak plasma concentrations of 1 to 10 µmol/L, whereas high-dose infusion regimens using 1.5 g/m2 or more yield peak levels of 0.1 to 1 mmol/L.251 Whether plasma concentrations are strictly proportional to dose is unclear. The initial phase of drug disappearance from plasma has a half-life of 2 to 3 hours, with no apparent variation as doses are increased to the high-dose range. This phase extends for the first 12 to 24 hours after drug administration and is largely determined by the rate of renal excretion of MTX. Prolongation of this phase, as well as of the terminal phase of drug disappearance from plasma, is observed in patients with renal dysfunction, in whom half- life is approximately proportional to the serum creatinine.259 Bressolle and colleagues260 studied the effects of moderate renal insufficiency on pharmacokinetics of MTX in patients with rheumatoid arthritis. Intramuscular MTX was administered to four separate groups of patients segregated according to creatinine clearance: less than 45, 45 to 60, 61 to 80, and more than 80 mL/minute. Noting an increased elimination half-life and reduced total clearance that correlated with the degree of renal impairment, the authors concluded that individual renal function testing was preferred over a general decrease of the MTX dose based only on observed serum creatinine. In patients with normal creatinine clearance, the half-life of the initial phase of drug disappearance increases with advancing age of the patient, which lends additional variability to plasma levels, disappearance kinetics, and toxicity.

The final phase of drug disappearance has a considerably longer half-life of 8 to 10 hours256, 258; this half-life may further lengthen in patients with renal dysfunction or with third-space fluid such as ascites. After conventional doses of 25 to 200 mg/m2, this terminal phase begins at drug concentrations above the threshold for toxicity to bone marrow and gastrointestinal epithelium. Thus, any prolongation of the terminal half-life is likely to be associated with significant toxicity.

The use of constant-infusion MTX has received increasing consideration because it offers the advantage of providing predictable blood and cerebrospinal fluid (CSF) concentrations for a specific period of time. Bleyer261 has used the following formulas for achieving a desired plasma concentration in patients with normal renal function:

An approximate correction for renal function may be made by reducing the infusion doses in proportion to the reduction in creatinine clearance, based on a normal creatinine clearance of 60 mL/minute per square meter. The terminal elimination phase of MTX increases from 3 to 5 hours in proportion to the duration of the constant infusion over the range of 24 to 72 hours.262 This variation probably represents slow tissue release of poorly effluxable MTX polyglutamates that form in cells in a time-dependent and dose-dependent manner.

The plasma pharmacokinetics of MTX may independently predict relapse in children treated during the maintenance phase of ALL with intermediate drug doses (1 g/m2). In a study of 108 children, a rapid drug clearance (84 to 132 mL/minute per square meter) was associated with a 40% risk of relapse, whereas those children with relatively slow drug clearance (45 to 72 mL/minute per square meter) had a significantly decreased (P2 = .01) risk of relapse (25%).263Steady-state MTX concentrations of less than 16 µmol/L were also associated with a lower probability of remaining in remission (P <.05) than in patients with concentrations in excess of 16 µmol/L.264 These associations were further supported by the finding of a systemic clearance of 123 mL/minute per square meter in 25 children who relapsed with ALL versus 72 mL/minute per square meter in 33 children who remained in continuous remission.265 These results suggest that dose should be increased in children with rapid drug clearance and low steady-state MTX levels.266

Renal Excretion

The bulk of drug is excreted in the urine in the first 12 hours after administration, with renal excretion varying from 44% to virtually 100% of the administered dose.258, 267 The higher figure is likely to be true for patients with normal renal function. MTX clearance by the kidney has exceeded creatinine clearance in some patients studied.268

During high-dose infusion, rapid drug excretion may lead to high MTX concentrations in the urine. These concentrations, approaching 10 mmol/L, exceed the solubility of the drug below pH 7.0 (Table 6.3) and are believed to be responsible for intrarenal precipitation of drug and renal failure. Thus, in high-dose regimens, hydration and alkalinization of the urine are recommended to avoid renal toxicity (Table 6.4). To ensure adequate intrarenal dissolution of MTX during high-dose therapy (0.7 to 8.4 g/m2), a 20-fold greater urine flow is required at pH 5.0 (2 to 42 mL/minute per square meter) than at pH 7.0 (0.1 to 1.2 mL/minute per square meter).269 Intensive hydration does not affect the clearance of MTX or the plasma pharmacokinetics, aside from its effects on the prevention of renal damage.270

The exact mechanism of MTX excretion in the human kidney has not been fully elucidated. In dog and monkey models, active secretion of MTX takes place in the proximal renal tubule, with reabsorption in the distal tubule.271 As noted earlier, the high clearance values in excess of creatinine clearance suggest that active tubular secretion of MTX occurs in humans. MTX excretion is inhibited by weak organic acids such as aspirin,215 piperacillin,272, 273 penicillin G,274oxacillin,275 ciprofloxacin,276 and probenecid, an inhibitor of organic acid secretion.272 The cephalosporins, including ceftriaxone, ceftriazone, ceftazidime and sulfamethoxazole, enhance the renal elimination of MTX, probably through competition for tubular reabsorption.273, 274 Simultaneous folic acid administration blocks MTX reabsorption, which suggests that leucovorin might accelerate MTX excretion in high-dose rescue regimens.



Solubility (mg/mL)

pH 5.0

pH 6.0

pH 7.0





7-Hydroxy methotrexate




2,4-diamino-N10-methyl pteroic acid




In high-dose MTX therapy, despite interindividual variation in pharmacokinetics, blood levels of drug may be accurately predicted by a preliminary determination of drug clearance using a small test dose (10 to 50 mg/m2).268, 277 Pharmacokinetic measurements made after delivery of this test dose provide a basis for calculating high-dose infusion rates according to the following formula (units of conversion must be carefully considered):

Thus, the desired infusion rate is the product of the target steady-state concentration multiplied by the MTX clearance rate, as determined from the test dose.268

Extrapolation from the test dose of 50 mg/m2 to a high-dose infusion schedule has proven to be reliable as long as renal function remains normal during the infusion period. The test dose technique has also been used to identify the subset of patients with impaired MTX elimination who are at increased risk for toxicity.278

Hepatic Uptake and Biliary Excretion

MTX undergoes uptake, storage, and metabolism in the liver, but the relative contribution of each of these processes to drug pharmacokinetics is unclear.

MTX is actively transported into hepatocytes by an uptake system that appears to have several components.279, 280 In the hepatocyte, MTX is converted to polyglutamate forms that persist for several months after drug administration.281 The parent drug also undergoes excretion into the biliary tract and is reabsorbed into the systemic circulation from the small intestine. Dactinomycin (actinomycin D) strongly inhibits biliary secretion of MTX, with little effect on hepatic uptake, and causes a marked increase in intrahepatic levels of the drug.282 Although this pharmacokinetic effect has not been examined in detail in humans; the possibility exists that the combination of MTX and dactinomycin may increase MTX levels in liver and enhance its hepatic toxicity.

The effects of conjugated and unconjugated bile salts on the enterohepatic circulation of MTX have been investigated in vivo in perfused rat intestinal preparations.283 This study demonstrated a saturable intestinal transport mechanism (Kt = 0.98 µmol/L). The unconjugated bile salt deoxycholate and the conjugated salt taurocholate significantly diminished MTX absorption. Folic acid, 5-methyltetrahydrofolate, and the organic anions rose bengal and sulfobromophthalein, also inhibited transport. These compounds might be useful in altering both systemic and specific hepatic toxicities associated with MTX by promoting MTX excretion in the stool.


Hydration and Urinary Alkalinization

Administer 2.5–3.5 L/m2/d of IV fluids starting 12 hr before and for 24–48 hr after administration of MTX drug infusion. Sodium bicarbonate 45–50 mEq/L IV fluid to ensure that urine pH is >7.0 at the time of drug infusion.

Commonly Used Drug Infusion Regimens

MTX Dose

Duration (hr)

Fluid (L/24 hr)

Bicarbonate (mEq/24 hr)

Leucovorin Calcium rescue

Onset of Rescue after Start of MTX)

1.5–7.5 g/m2




15 mg IV q3hr × then 15 mg PO q6hr × 7


8–12 g/m2




10 mg PO q6hr × 10


3.0–7.5 g/m2




10 mg/m2 IV × 1, then 10 mg/m2 PO q6hr × 12


1 g/m2




15 mg/m2 IV q6hr × 2, then 3 mg/m2 PO q12hr × 3


1.0–7.5 g/m2




10 mg/m2 IV × 1, then 10 mg/m2 PO q6hr × 11


IV, intravenously; NS, not specified; PO, per os.
Adapted from Ackland SP, Schilsky RL. High-dose methotrexate: a critical reappraisal. J Clin Oncol 1987;5:2017–2031.

Monitor Points
MTX drug levels above 5 × 10-7 mol/L at 48 hr after the start of MTX infusion require continued leucovorin rescue.a A general guideline for leucovorin rescue is as follows:

MTX Level

Leucovorin Dosage

5 × 10-7mol/L

15 mg/m2 q6h × 8

1 × 10-6mol/L

100 mg/m2 q6h × 8

2 × 10-6mol/L

200 mg/m2 q6h × 8

aMTX drug levels should be measured every 24 hr and the dosage of leucovorin adjusted until the MTX level is <5 × 10-8 mol/L.


Widely divergent estimates have been made of the relative importance of biliary excretion of MTX in humans. Lerne et al.,284 using tritium-labeled MTX, found only 0.41% of an administered dose in the bile of a patient with a biliary fistula. Subsequent studies have variously estimated that 6.7 to 9%285 or 20%267 of an administered dose enters the biliary tract. Calvert and co-workers267 used the highly specific DHFR inhibition assay to measure biliary concentrations of MTX and found that biliary levels were 2,500-fold to 10,000-fold higher than simultaneous concentrations in the plasma. Despite these high concentrations in bile, less than 10% of an intravenous dose of MTX is eliminated in the feces. In the presence of diminished renal function, the enterohepatic circulation may become an important determinant of drug elimination.286 Under these conditions, intestinal binding to prevent reabsorption of the drug with activated charcoal287 or the anion-exchange resin cholestyramine,288 which has a 5.4-fold greater binding capacity than charcoal, may be used to enhance the nonrenal excretion of MTX.


The introduction of high-dose MTX regimens has led to the identification of at least two MTX metabolites in humans. Jacobs et al.289 identified 7-OH-MTX in the urine of patients receiving high-dose infusions; 7-OH-MTX constituted 20 to 46% of material excreted in the urine in the interval between 12 and 24 hours after the start of the infusion. The fraction of drug in the form of this metabolite was estimated to be as high as 86% in the period from 24 to 48 hours. A second metabolite, DAMPA, has also been identified in plasma and urine, and, at later times, this metabolite makes up an important fraction of drug-derived material, comprising a mean of 25% of material excreted in the interval from 24 to 48 hours.241 Both of these metabolites are known to accumulate in plasma, and at 24 to 48 hours after high-dose MTX administration, they account for most of the MTX-derived material found in plasma and may give spuriously high MTX values when using certain assays

Regarding the sites of MTX metabolism, 7-OH-MTX is probably formed through the action of the liver enzyme aldehyde oxidase where levels of the 7-OH-MTX metabolite have been found to be 700-fold higher in bile than in serum.290 The extent to which polyglutamated 7-OH-MTX is formed in malignant but not in normal cells may be important in the selective action of MTX because the 7-OH polyglutamates have measurable effects on several critical intracellular pathways. The polyglutamates of 7-OH-MTX are inhibitors of the folate-dependent enzymes AICAR transformylase and TS, with potency similar to that of MTX polyglutamates (Ki = 3 and 0.4 µmol/L, respectively).291 These metabolites can bind to DHFR109,153 but are relatively weak inhibitors (Ki = 9 nmol/L) of this enzyme compared with MTX.165, 175, 292 Unlike with the inhibition of certain folate-dependent enzymes, polyglutamation has relatively minor effects on the potency of DHFR inhibition by 7-OH-MTX.165, 292 Exposure of murine leukemia cells to high concentrations of 7-OH-MTX (100 µmol/L) resulted in only mild inhibition of cell growth.208 Despite primary metabolism of MTX to the 7-OH-MTX metabolite by the liver, no dosage adjustment of MTX appears to be necessary for patients with hepatic dysfunction.

The pteroic acid metabolite DAMPA is probably formed by the action of bacterial carboxypeptidases in the gastrointestinal tract. Enzymes specific for glutamate terminal peptide bonds have been characterized.293 DAMPA is also produced by the enzymatic cleavage of MTX in a rescue regimen initially tested in the treatment of brain tumors.294 In this protocol, high-dose systemic MTX is followed by the infusion of the bacterial enzyme carboxypeptidase G1, which degrades MTX in the systemic circulation but leaves drug intact in the brain and CSF.

The role of these metabolites in producing MTX toxicity or influencing therapeutic activity is uncertain. Both 7-OH-MTX and the pteroic acid metabolite are less soluble than the parent drug (Table 6.3). Jacobs and colleagues289 demonstrated that 7-OH-MTX constituted more than 50% of precipitated intrarenal material in their study of MTX-induced renal failure in monkeys, but the role of either metabolite in the clinical syndrome of MTX-associated nephrotoxicity in humans is unproven.


Primary Toxic Effects

The primary toxic effects of folate antagonists are myelosuppression and gastrointestinal mucositis. The incidence of these and other toxicities depends on the specific dose, schedule, and route of drug administration and is summarized in Table 6.5. The intestinal and oral epithelia are somewhat more sensitive than granulocyte and platelet precursors in that drug schedules that produce intense mucositis (particularly those with prolonged, low drug concentrations) may cause little marrow suppression. The threshold plasma concentration of MTX required to inhibit DNA synthesis in bone marrow has been estimated to be 10 nmol/L, whereas gastrointestinal epithelium is inhibited at 5 nmol/L plasma concentrations.295 This greater sensitivity of gastrointestinal epithelium is believed to result from greater accumulation and persistence of MTX in intestinal epithelium than in bone marrow.296 Mucositis usually appears 3 to 7 days after drug administration and precedes the onset of a fall in white blood count or platelet count by several days. In patients with compromised renal function, small doses on the order of 25 mg may provide cytotoxic blood levels for up to 3 to 5 days and may result in serious bone marrow toxicity. Myelosuppression and mucositis usually are completely reversed within 2 weeks, unless drug excretion mechanisms are severely impaired.

Methylenetetrahydrofolate reductase (MTHFR) is a critical enzyme in folate metabolism that regulates the metabolism of folate, methionine and homocysteine. A C-to-T polymorphism of this enzyme at amino acid residue 677 results in a marked reduction in the activity of the enzyme particularly in individuals homozygous for this polymorphism. Multiple clinical studies in patients undergoing therapy with MTX for the management of rheumatoid arthritis, leukemia, ovarian cancer, and in the setting of bone marrow transplantation have reported a consistently elevated level of homocysteine following MTX exposure and a marked increase in the level of toxicity associated with MTX in patients homozygous for the 677C to T polymorphism.297, 298, 299, 300, 301 These studies suggest that either assessment for the TT genotype or the occurrence of marked elevations in serum hymocycteine levels following MTX exposure may be useful as markers for predicting high-grade MTX toxicity.

The introduction of high-dose MTX regimens with leucovorin rescue302 has been associated with a spectrum of clinical toxicities that has required more careful monitoring of drug pharmacokinetics in individual patients. These regimens use otherwise lethal doses in a 6- to 36-hour infusion, followed by a 24- to 48-hour period of multiple leucovorin doses to terminate the toxic effect of MTX. Several of the more commonly used high-dose regimens and their related pharmacokinetics are presented in Table 6.4. For each regimen, successful rescue by leucovorin depends on the rapid elimination of MTX by the kidneys. Early experience with high-dose regimens, however, indicated that MTX itself may have acute toxic effects on renal function during the period of drug infusion, which can lead to delayed drug clearance, ineffective rescue by leucovorin, and a host of secondary toxicities, including severe myelosuppression, mucositis, and epithelial desquamation.258 During the early clinical trials of high-dose MTX, a number of toxic deaths were recorded.303

The cause of drug-induced renal dysfunction, which is usually manifested as an abrupt rise in serum blood urea nitrogen and creatinine with a corresponding fall in urine output, is thought to arise from the precipitation of MTX and possibly its less soluble metabolites, 7-OH-MTX and DAMPA, in acidic urine.289 A direct toxic effect of antifolates on the renal tubule, however, has been suggested by the observation that aminopterin, an equally soluble compound that is used at one-tenth the dose of MTX, is also associated with renal toxicity; however, a direct nephrotoxic role of MTX has not been substantiated in clinical investigations.304 Jacobs et al.289 were able to reproduce the syndrome of MTX-induced renal failure in a monkey model system and demonstrated precipitation of both MTX and 7-OH-MTX in the renal tubules. Both of these compounds have limited solubility under acid pH conditions. To prevent precipitation, most centers use vigorous hydration (2.5 to 3.5 L of fluid per square meter per 24 hours, beginning 12 hours before MTX infusion and continuing for 24 to 48 hours), with alkalinization of the urine (45 to 50 mEq of sodium bicarbonate per liter of intravenous fluid). The MTX infusion should not begin until urine flow exceeds 100 mL/hour and urine pH is 7.0 or higher, and these parameters should be carefully monitored during the course of drug infusion (Table 6.4).







Pulmonary Toxicity


Intermediate IV (50–100 mg/m2)



+ (transaminasemia)




High-dose IV with leucovorin calcium (100–12,000 mg/m2)


+++ (requires urinary alkalinization and hydration)

++ (transaminasemia)



++ (acute and chronic)

Low-dose PO, daily dose (5–25 mg/m2)



+++ (up to 25% cirrhosis)




Low-dose PO, pulse therapy (5–25 mg/m2)



++ (rarely cirrhosis)










++ (acute, subacute, and chronic)

+, some toxicity; ++, moderate toxicity; +++, high toxicity; -, no toxicity; ±, possible toxicity.
IV, intravenously; PO, per os.

With this regimen, the incidence of renal failure and myelosuppression has been markedly reduced. No change in the rate of MTX excretion or alteration of plasma pharmacokinetics results from the intense hydration used in the preparatory regimen previously described270; thus, these safety measures should have no deleterious effect on the therapeutic efficacy of the regimen. Despite careful attention to the details of hydration and alkalinization, occasional patients can develop serious or even fatal toxicity.303 Almost all of these toxic episodes are associated with delayed MTX clearance from plasma and can be predicted by routine monitoring of drug concentration in plasma at appropriate times after drug infusion.305 In an analysis of 790 patients treated with high-dose MTX for osteosarcoma, the incidence of delayed MTX clearance (>5umol/L at 24 hours postinfusion) was 1.6% per cycle of treatment.306 The specific time for monitoring, and the guidelines for distinguishing between normal and dangerously elevated levels, must be determined for each regimen and for each assay procedure. In general, a time point well into the final phase of drug disappearance, such as 24 or 48 hours after the start of infusion, should be chosen (Table 6.4). The use of ketoprofen and other nonsteroidal anti-inflammatory drugs (NSAIDs) has been associated with severe MTX toxicity.307 A review of 118 cases of single-agent, high-dose MTX therapy revealed four cases of fatal toxicity associated with the use of NSAIDs. Patients treated with NSAIDs demonstrated a marked prolongation of serum MTX half-life that was postulated to be caused by decreased renal elimination secondary to inhibition of renal prostaglandin synthesis or competitive inhibition of human organic anion transporters (hOAT) responsible for MTX uptake in the kidney.308, 309

Early detection of elevated concentrations of MTX allows institution of specific clinical measures. Continuous medical supervision is warranted until the severity and duration of myelosuppression can be determined. Leucovorin in increased doses is required and must be continued until plasma MTX concentration falls below 50 nmol/L. Because of the competitive relationship between MTX and leucovorin, the leucovorin dose must be increased in proportion to the plasma concentration of MTX. Small doses of leucovorin are unable to prevent toxicity in patients with elevated drug levels, even when leucovorin is continued beyond 48 hours.305 As a general rule, a reasonable course is to treat with leucovorin at a dosage of 100 mg/m2 every 6 hours for patients with MTX levels of 1 µmol/L and to increase this dosage in proportion to the MTX level up to a maximum of 500 mg/m2 (Table 6.4). Subsequent leucovorin dosage adjustments should be based on repeated plasma MTX levels taken at 24-hour intervals. The results of in vitro studies indicate that leucovorin alone may not be able to rescue patients with plasma MTX concentrations above 10 µmol/L.

The absorption of oral leucovorin is saturable such that the bioavailability of the compound is limited above total doses of 40 mg. The fractional absorption of a 40-mg dose is 0.78, whereas that of 60- and 100-mg doses is 0.62 and 0.42, respectively. For this reason, leucovorin is usually administered intravenously to assure its absorption.

Because of the variable effectiveness of leucovorin in preventing toxicity in patients with levels of 10 µmol/L or greater at 48 hours, alternative methods of rescue have been proposed. Both intermittent hemodialysis and peritoneal dialysis are ineffective in removing significant quantities of MTX. However, continuous flow hemodialysis has been effective in reducing plasma MTX concentration and preventing toxicity in patients with MTX-induced failure.309a The use of charcoal hemoperfusion columns is capable of removing MTX and other antineoplastic drugs from whole blood and has been applied successfully in a few patients; however, platelet adherence to these columns may lead to thrombocytopenia.

Abelson and co-workers294 used a bacterial enzyme, carboxypeptidase G1, which inactivates MTX by removal of its terminal glutamate, to destroy circulating MTX. The regimen of high-dose MTX followed by intravenous carboxypeptidase was well tolerated, but this form of enzymatic rescue carries a risk of hypersensitivity to the bacterial enzyme. Bertino et al.310 have demonstrated the feasibility of attaching the enzyme to hollow fiber tubing, which can then be used in an extracorporeal shunt for drug removal and thus avoid immune sensitization. One potential disadvantage of carboxypeptidase G1, however, is its relatively high affinity for natural folates as well as MTX. DeAngelis and colleagues311 conducted a pilot study to determine the efficacy of carboxypeptidase G2 (CPG2) rescue after high-dose MTX in patients with recurrent cerebral lymphoma. All patients had at least a 2-log decline in plasma MTX levels within 5 minutes of CPG2 administration, whereas CSF MTX concentrations remained elevated for 4 hours after CPG2. No MTX or CPG2 toxicity was observed, and anti-CPG2 activity antibodies were not detected in any patient. The authors concluded that CPG2 rescue was a safe and effective alternative to leucovorin rescue after high-dose MTX chemotherapy. Additional utility of CPG2 as a safe and effective means for preventing severe MTX toxicity was demonstrated in several reports of patients with delayed MTX clearance.312, 313 In an NCI study on the use of CPG2 in pediatric patients who developed nephrotoxicity while receiving high-dose MTX, Widemann et al.314 found that CPG2 and thymidine rescue was well tolerated and resulted in a rapid and effective reduction in the plasma MTX concentration with only mild-to-moderate MTX-related toxicity.

Preliminary reports have described the successful prevention of MTX toxicity in humans using 6- to 40-hour infusions of the antifolate followed by a 72-hour infusion of thymidine at a rate of 8 g/m2 per day.229, 315, 316 Patients receiving a bolus dose of MTX, 3 g/m2, were successfully rescued by thymidine infusion (1 g/m2 per day); the infusion was begun 24 hours after MTX administration and continued until MTX concentration in the plasma reached 50 nmol/L.317

In the experimental setting, MTX toxicity can also be blocked by drugs that prevent cell progression into the S phase of the cell cycle. The antagonistic effect of L-asparaginase on MTX toxicity is a representative example; through depletion of the amino acid asparagine, L-asparaginase, inhibits protein synthesis and prevents entry of cells into the DNA-synthetic phase of the cell cycle.318 Rescue regimens319 that use high doses of MTX (up to 400 mg/m2), followed within 24 hours by 20,000 to 40,000 U per m2 of L-asparaginase, produce minimal bone marrow toxicity and mucositis, and appear to have some effectiveness in patients refractory to low-dose MTX alone. Yap et al.319 reported a complete remission rate of 62% (13 of 21 cases) in adult patients with ALL who had failed initial therapy with conventional induction regimens.

Drugs that inhibit the TS reaction, thereby preventing alterations in the composition of the intracellular folate pools and negating the effect of DHFR enzyme inhibition, also prevent MTX toxicity. This effect has been studied in detail and seems to explain the antagonism of fluoropyrimidine pretreatment followed by MTX.320

Poor nutritional status has been associated with an increased risk of toxicity from MTX.321, 322 Poorly nourished patients appear to have an approximately twofold decrease in their clearance of MTX. The reason for this delayed drug clearance appears to be a protracted enterohepatic circulation. Providing dietary protein in the form of polypeptides (rather than amino acids) either alone or in combination with cholestyramine treatment to bind intestinal MTX may be useful in avoiding excess toxicity associated with nutritional deficiencies.

Other Toxicities


In addition to its inhibitory effects on rapidly dividing tissues, MTX has toxic effects on nondividing tissues not easily explained by its primary action on DNA synthesis. Long-term MTX therapy is associated with portal fibrosis, which may, on occasion, progress to frank cirrhosis. Chronic liver disease has occurred most frequently in patients with psoriasis or rheumatoid arthritis or in children with acute leukemia who have received maintenance therapy over a period of several years. The incidence of cirrhosis has been estimated to be 10% in MTX-treated patients with psoriasis but may reach as high as 25 to 30% in those patients treated for 5 years or longer with continuous daily therapy.323 Cirrhosis does not always progress with continued antifolate treatment. Of 11 patients with psoriasis who showed cirrhotic changes on liver biopsy and continued to receive treatment, only 3 showed progression on subsequent biopsy, and 3 had no pathologic findings on a follow-up biopsy.323 The use of “pulsed” weekly therapy rather than continuous daily treatment appears to lessen the incidence of MTX-associated hepatotoxicity.324, 325 Several studies suggest that the incidence of hepatic cirrhosis is no different in patients with rheumatoid arthritis treated with MTX pulse therapy than in untreated patients despite long-term therapy (longer than 5 years).326, 327 Evidence of hepatic toxicity was detected on liver biopsy in 76% of 29 patients receiving weekly (7.5-mg) pulse therapy with MTX for rheumatoid arthritis; however, only 1 patient had severe fibrosis.326 These patients had been treated for an average of approximately 2.5 years (1,500 mg total dose). Abnormal elevations in serum transaminases have been found in up to 70% of patients treated with long-term weekly MTX; however, the enzyme elevations were poor predictors of liver damage.328 In three series of patients who underwent liver biopsy while being treated with long-term weekly MTX, 17 to 30% had fibrotic changes but 0 to 3% demonstrated cirrhosis.328, 329

Acute elevations of liver enzymes are commonly observed after high-dose MTX administration and usually return to normal within 10 days. The frequency and severity of liver enzyme elevations appear to be directly related to the number of MTX doses received.330 Liver biopsy in such patients has revealed fatty infiltration but no evidence of hepatocellular necrosis or periportal fibrosis. The late occurrence of cirrhosis in patients treated with high-dose MTX has not been reported.


Treatment with MTX is associated with a poorly characterized, self-limited pneumonitis, with fever, cough, and an interstitial pulmonary infiltrate.331, 332Eosinophilia has not been a consistent finding, either in the peripheral blood or in open lung biopsy specimens. Lung biopsies have revealed a variety of findings, from simple interstitial edema and a mononuclear infiltrate to noncaseating granulomas. The possibility that MTX pneumonitis may not represent a hypersensitivity phenomenon has been raised because of the failure of some patients to react to reinstitution of MTX therapy. However, bronchoalveolar lavage in three patients with presumptive MTX-induced lung damage revealed a predominance of T8 suppressor lymphocytes. In contrast to peripheral lymphocytes obtained from MTX-treated patients with no lung damage, lymphocytes from the study patients elaborated leukocyte inhibitory factor in response to MTX exposure.333 This study supports an immunologic basis for MTX-related lung damage. The possibility exists, however, that many case reports of “MTX lung” in fact represent unrecognized viral infections or allergic reactions to unsuspected allergens. With the use of long-term weekly low-dose MTX therapy for rheumatoid arthritis, however, a number of cases of MTX-associated lung damage have been reported.334, 335 A review of 168 patients treated with MTX for rheumatoid arthritis uncovered 9 cases (5%) of probable MTX-associated lung toxicity.336 Using a retrospective combined-cohort review and abstraction from the medical literature, Kremer and colleagues337 characterized the clinical features of MTX-associated lung injury in patients with rheumatoid arthritis. Clinical symptoms of MTX toxicity in the cohort included the subacute development of shortness of breath (93%), cough (82%), and fever (69%), with resultant death in 5 of 27 patients. The authors concluded that early symptom recognition and the cessation of MTX administration could avoid the serious and sometimes fatal outcome of this MTX-associated toxicity. Corticosteroids have been used in a small number of patients who ultimately recovered,338 but the utility of this approach has yet to be established. Alarcon et al.339 found that the strongest predictors of MTX-induced lung injury in rheumatoid arthritis patients included older age, presence of diabetes, rheumatoid pleuropulmonary involvement, presence of hypoalbuminemia, and prior use of disease-modifying antirheumatic drugs.


True anaphylactic reactions to MTX are rare. Two cases of acute hypersensitivity reaction to MTX have been described.340 In the first case, the patient experienced acute cardiovascular collapse, which was reproduced on rechallenge of the patient with MTX. In the second case, the acute reaction consisted of facial edema, rash, and generalized pruritus, and again was elicited on rechallenge. Both patients were receiving bacille Calmette-Guérin in conjunction with MTX at the time of these reactions, and thus may have developed a heightened sensitivity to MTX. Three cases of toxic erythema and desquamation of the hands were reported in patients receiving high doses (1.5 g/m2) of MTX for the treatment of non-Hodgkin's lymphoma.341 This toxic reaction was associated with severe mucositis and was ameliorated by MTX dose reductions on subsequent treatment.

Reversible oligospermia with testicular failure has been reported in men treated with high-dose MTX.342 No alterations in follicle-stimulating hormone, luteinizing hormone, estradiol, or progesterone have been observed in women exposed to MTX.


Because of its high degree of ionization at physiologic pH, MTX penetrates into the CSF with difficulty. During a constant intravenous drug infusion,343 the ratio of venous MTX concentration to CSF concentration is approximately 30:1 at equilibrium. Thus, plasma levels in excess of 30 µmol/L would be required to achieve the concentration of 1 µmol/L that is thought to be necessary for killing of leukemic cells. Protocols for prophylaxis against meningeal leukemia and lymphoma using systemic high-dose infusions of MTX have demonstrated that high-dose MTX infusions are a reasonable treatment alternative to intrathecal prophylaxis. Overt meningeal leukemia increases the CSF: plasma ratio and experience supports the use of MTX at a loading dose of 700 mg/m2 followed by a 23-hour infusion of 2,800 mg/m2, with leucovorin rescue as an excellent treatment alternative for patients with carcinomatous meningitis capable of achieving the requisite CSF levels of 1 µmol/L.344, 345 In children with ALL, a diminished CSF: plasma ratio has been found to be a useful predictor of CNS relapse.346

Direct intrathecal injection of MTX has been used for the treatment and prophylaxis of meningeal malignancy. The readers are referred to a comprehensive review on this topic.347 Drug injected into the intrathecal space distributes in a total volume of approximately 120 mL for patients over 3 years of age. Thus, a maximal total dose of 12 mg is advised for all patients over 3 years, with lower doses indicated for younger children. Bleyer261 has recommended a dose of 6 mg for age 1 or younger, 8 mg for ages 1 to 2, and 10 mg for ages 2 to 3. The peak CSF concentration achieved by this schedule is approximately 100 µmol/L. Lumbar CSF drug concentrations decline in a biphasic pattern with a terminal half-life of 7 to 16 hours. This terminal phase of disappearance may be considerably prolonged in patients with active meningeal disease and in older-age patients.347, 348 Injection of radiolabeled MTX into the ventricular space of rabbits demonstrated rapid but variable distribution of MTX in the gray matter adjacent to the CSF, which suggests a mechanism for the various syndromes associated with MTX neurotoxicity.349 MTX is cleared from spinal fluid by bulk resorption of spinal fluid (i.e., “bulk flow”), a process that may be prolonged by increases in intracranial pressure and the administration of acetazolamide. A second component of resorption involves the active transport of this organic anion by the choroid plexus. A prolongation of the terminal half-life is also found in patients who develop drug-related neurotoxicity, although a causal relationship between abnormal pharmacokinetics and neurotoxicity has not been firmly established.

MTX administered into the lumbar space distributes poorly over the cerebral convexities and into the ventricular spaces.343 The concentration gradient between lumbar and ventricular CSF may exceed 10:1. Although this uneven distribution has no documented role in determining clinical relapse of patients treated for meningeal leukemia, awareness of this potential problem has led to clinical trials using direct intraventricular injection of MTX via an Ommaya reservoir. Bleyer and colleagues350 have demonstrated that a concentration × time regimen in which 1 mg MTX was injected into the Ommaya reservoir every 12 hours for 3 days yielded continuous CSF levels above 0.5 µmol/L and achieved therapeutic results equivalent to those with the conventional intralumbar injection of 12 mg every 4 days. Moreover, this concentration × time regimen was associated with a considerable reduction in neurotoxic side effects, presumably owing to the avoidance of high peak levels of drug associated with higher MTX doses. Glantz et al.351 reported on the use of high-dose intravenous MTX as the sole treatment for nonleukemic meningitis. Sixteen patients with solid tumor neoplastic meningitis received high-dose intravenous MTX (8 g/m2over 4 hours) with leucovorin rescue. Compared with a reference group of patients receiving standard intrathecal MTX, the high-dose intravenous group exhibited cytotoxic CSF and serum MTX concentrations that were maintained much longer than with intrathecal dosing. In addition, median survival in the high-dose intravenous MTX group was 13.8 months versus 2.3 months for the intrathecal reference group (P = .003).

Three different neurotoxic syndromes have been observed after treatment with intrathecal MTX.350 The most common and most immediate neurotoxic side effect is an acute chemical arachnoiditis manifested as severe headache, nuchal rigidity, vomiting, fever, and inflammatory cell pleocytosis of the spinal fluid. This constellation of symptoms appears to be a function of the frequency and dose of drug administered, and may be ameliorated either by reduction in dose or by a change in therapy to intrathecal cytosine arabinoside. A less acute but more serious neurotoxic syndrome has been observed in approximately 10% of patients treated with intrathecal MTX. This subacute toxicity appears during the second or third week of treatment, usually in adult patients with active meningeal leukemia, and is manifested as motor paralysis of the extremities, cranial nerve palsy, seizures, or coma. Because MTX pharmacokinetics is abnormal in these patients, the suspicion is that this subacute neurotoxicity may be the result of extended exposure to toxic drug concentrations.347 Finally, a more chronic demyelinating encephalopathy has been observed in children months or years after intrathecal MTX therapy. The primary symptoms of this toxicity are dementia, limb spasticity, and, in more advanced cases, coma. Computerized axial tomography (CT) has revealed ventricular enlargement, white matter changes, cortical thinning, and diffuse intracerebral calcification in children who have received prophylactic intrathecal MTX.352, 353 Most of these patients had also received cranial irradiation (>2,000 rad) and all had received systemic chemotherapy.

Treatment with repeated courses of high-dose intravenous MTX may also result in encephalopathy.354 In these patients, symptoms of dementia and paresis may develop in the second or third month after treatment and may also be associated with diffuse cortical hypodensities on CT scan. A second form of cerebral dysfunction associated with high-dose MTX is an acute transient dysfunction, which has been described in 4 to 15% of treated patients.355, 356, 357 The syndrome consists of any combination of paresis, aphasia, behavioral abnormalities, and seizures. The neurologic events occur an average of 6 days after the MTX dose and completely resolve, usually within 48 to 72 hours.

Patients may have received any number of MTX doses before the onset of this neurotoxic event, and some patients may have repeat episodes with subsequent MTX doses. In general, CSF and head CT scans are normal, but low-density lesions have been noted in some cases.358 The electroencephalogram may represent the only abnormal study and shows a diffuse or focal slowing. No clinical evidence exists to support the use of leucovorin, either acutely after intrathecal MTX or over the long term in patients who develop neurotoxic symptoms. Although leucovorin can enter the CSF, its penetration appears to be poor.359, 360 A comparison of neurologic toxicities was undertaken in a randomized trial involving 49 children with acute leukemia treated with either intrathecal MTX plus radiation or high-dose systemic MTX for central nervous system prophylaxis.361 Long-term toxicities were similar with either treatment option, and overall decreases in intelligence quotients were found to be clinically significant in 61% of the children. In addition, 58% of the patients treated with systemic therapy had abnormal electroencephalograms and 57% of those treated with intrathecal MTX and radiation experienced somnolence syndrome. Mahoney and colleagues362 described the incidence of acute neurotoxicity in 1,304 children with lower risk B-precursor lymphoid leukemia treated as part of the Pediatric Oncology Group trial. After remission induction, patients were randomized into one of three 24-week intensification schedules (intermediate-dose MTX or divided-dose oral MTX with or without intravenous mercaptopurine and extended intrathecal therapy). Overall, acute neurotoxicity occurred in 7.8% (95 of 1,218) of eligible patients, and the authors found that intensification with repeated intravenous MTX and low-dose leucovorin rescue was associated with a higher risk of acute neurotoxicity and leukoencephalopathy, especially in patients who received concomitant triple intrathecal therapy (MTX, dexamethasone, and cytosine arabinoside).

The etiology of the MTX-associated neurotoxicity is unknown. Vascular events in the form of vasospasm or emboli have been proposed to explain these neurologic abnormalities, and studies have suggested alterations in brain glucose metabolism after MTX treatment.363 Investigators from St. Jude Children's Research Hospital found that the incidence of seizures in children treated for acute leukemia with MTX were related to acute elevations in serum homocysteine levels following MTX treatment.364 Of interest, these investigators did not find an association between seizures and MTHFR genotype. Long-term exposure of rat cerebellar explants to 1 µmol/L MTX resulted in axonal death 2 weeks after drug exposure and loss of myelin sheaths in 5 weeks, which suggests a direct toxic effect of MTX on axonal cells.365 DHFR is present in brain tissue, but its biochemical role in the cerebral cortex, the primary site of MTX neurotoxicity, is uncertain. Several studies have demonstrated the ability of cranial radiation to increase blood-brain barrier permeability to serum proteins and MTX.366Because radiation and MTX are frequently used together, this interaction may be an important mechanism for enhanced toxicity. Inadvertent overdose of intrathecal MTX generally has a fatal outcome. Immediate lumbar puncture with CSF removal along with ventriculolumbar perfusion has been successfully used to avert catastrophe in such situations.367


A variety of dosage schedules and routes of administration are used clinically, including high-dose therapy with the addition of leucovorin rescue. The selection of an appropriate schedule depends largely on the specific disease being treated, on other antineoplastic agents or radiation to be used in combination regimens, on the patient's tolerance for host toxicity, and on other factors that might alter pharmacokinetics. Parenteral schedules are preferred for induction therapy regimens in which maximal concentrations and duration of exposure are desirable in an effort to achieve complete remission. High-dose MTX regimens and leucovorin rescue offer the advantage of minimal bone marrow toxicity. This regimen, however, can safely be used only in patients with normal renal and hepatic function and under conditions in which no large extracellular accumulations of fluid are present. As emphasized previously, high-dose regimens should be instituted only when plasma monitoring is available to determine the adequacy of drug clearance and the risk of serious toxicity. Furthermore, because leucovorin may rescue tumor cells as well as normal cells, the optimal dose, schedule, and clinical utility of high-dose MTX with leucovorin rescue needs to be more carefully defined.



Thymidylate synthase (TS) represents a logical target for new drug development using folate analogs, and pemetrexed (LY231514, Alimta), a pyrrolo(2,3-d)pyrimidine-based antifolate analog, is a potent inhibitor of TS. Pemetrexed is avidly transported into cells via the reduced-folate carrier (RFC) and possibly by a unique transporter identified in mesothelioma cell lines.368, 368a It is metabolized to the polyglutamated forms, which are potent inhibitors of several folate-dependent enzymatic reactions. The multitargeting effect of pemetrexed was seen in studies by Shih et al.368, who suggested that, at higher concentrations, pemetrexed and its polyglutamates not only act as TS inhibitors, depleting dTTP pools, but also inhibit other key folate-requiring enzymes, including glycinamide ribonucleotide formyltransferase, and to a lesser extent DHFR, 5-aminoimidazole-4-carboxamide ribonucleotide formyltransferase, and C1-tetrahydrofolate synthase. The combined inhibitory effects of pemetrexed give rise to a cellular level end-product reversal pattern that is different from those of other inhibitors such as MTX and the quinazoline antifolates. In addition, pemetrexed has less effect on the folate and nucleotide pools as compared with MTX.369 In studies evaluating the effects of folic acid on modulating the toxicity and antitumor efficacy of pemetrexed in human tumor cell lines adapted to growth in low-folate medium, folic acid was shown to be 100- to 1,000-fold less active than folinic acid at protecting cells from pemetrexed-induced cytotoxicity.370 Further, folic acid supplementation was demonstrated to preserve the antitumor activity of pemetrexed while reducing toxicity in mice. In patients with mesothelioma, pemetrexed as a single agent produced a response rate of 14%. In patients, severe toxicity was correlated with high serum concentrations of hemocysteine, an indicator of folate deficiency. In an additional 15% of patients, B12 deficiency was thought to be responsible for pemetrexed toxicity. Of interest, the addition of vitamin B12 (1 mg intramuscularly) and folic acid (1mg/day for 2 weeks beginning 2 weeks prior to pemetrexed) resulted in an improved toxicity profile and improved efficacy in patients with mesothelioma treated with pemetrexed.371, 372 In a randomized phase III investigation of 456 patients with mesothelioma, comparing cisplatin with the combination of cisplatin plus pemetrexed, the patients treated with the doublet enjoyed a significantly improved response rate (41 versus 17%), time to disease progression (5.7 versus 3.9 months), and overall survival (12.1 versus 9.3 months) when compared with single-agent cisplatin therapy. Pemetrexed has been demonstrated to have activity in several solid tumors including non–small cell lung cancer, in which it was shown to have a 16% response rate in untreated patients.373 A phase III trial with 571 patients randomized patients with advanced and refractory non–small cell lung cancer to either pemetrexed or docetaxel and showed that both drugs resulted in identical efficacy outcomes, but the pemetrexed was associated with a significantly improved toxicity profile.374 Aside from myelosuppression and intestinal toxicity, pemetrexed causes a rash in 40% of patients. The rash is suppressed by the administration of dexamethasome, 4 mg 2 times a day, on days 1, 0, and +1.


Nolatrexed (AG337, THYMITAQ) is a nonclassic inhibitor of TS specifically designed to avoid potential resistance mechanisms that can limit the activity of the classic antifolate antimetabolites.375 Nolatrexed is a lipophilic molecule designed using x-ray structure-based methods to interact at the folate cofactor binding site of the TS enzyme. TS was suggested as the locus of action of nolatrexed by the ability of thymidine to antagonize cell growth inhibition and the direct demonstration of TS inhibition in whole cells using a tritium-release assay.376 Nolatrexed is characterized as a non–glutamate-containing molecule that does not require facilitated transport for uptake and does not undergo, nor require, intracellular polyglutamylation for activity. L1210 cells treated with nolatrexed exhibited S-phase cell-cycle arrest and a pattern of nucleotide pool modulations, including a reduction in thymidine triphosphate levels, consistent with inhibition of TS.376 Rafi et al.375 measured plasma concentrations of deoxyuridine (dUrd) in patients receiving doses of nolatrexed at levels of more than 600 mg/m2 and found elevation in plasma dUrd levels (60 to 290%), which implied that TS inhibition was being achieved in patients. In all cases, dUrd concentrations quickly returned to pretreatment levels after the end of the infusion; this suggested that TS inhibition was not maintained, presumably because of the brief intracellular half-life of the nonpolyglutamated parent compound. A phase I trial evaluating intravenous administration of nolatrexed found dose-limiting myelosuppression and a high incidence of thrombotic phenomena.377 A second trial evaluating 5-day oral administration of nolatrexed showed rapid absorption with a median bioavailability of 89%; dose-limiting toxicities were gastrointestinal. The authors concluded that nolatrexed could be safely administered as an oral preparation at a dosage of 725 mg/m2 per day for 5 days.378 Using this dose and schedule, in 139 untreated patients with head and neck cancer, nolatrexed was found to have similar activity as MTX.379


Raltitrexed (ZD1694, Tomudex) is a water-soluble TS inhibitor that appears to have an acceptable toxicity profile, convenient dosing schedule, and antitumor activity in colorectal, breast, pancreatic, and a variety of other solid cancers.380, 381, 382, 383 This drug is a second-generation agent designed to overcome the major toxicity associated with its predecessor, CB3717, namely, poorly predictable nephrotoxicity. Yin and colleagues384 conducted in vitro studies on the human A253 head and neck squamous carcinoma cell line to evaluate the downstream molecular alterations induced by the potent and sustained inhibition of TS by raltitrexed. TS inhibition by raltitrexed resulted in a time-dependent induction of megabase DNA fragmentation followed by a secondary 50- to 300-kilobase DNA fragmentation, which may correlate with reduced expression of p27 and increase in cyclin E and cdk2 kinase activity. Cunningham382 reviewed the results of three large controlled studies that suggest that raltitrexed is an effective alternative to 5-fluorouracil–based therapy in patients with advanced colorectal cancer, and that raltitrexed has the advantage of a predictable toxicity profile, minimization or avoidance of mucositis, and convenient dosing schedule. The data concerning progression-free survival and survival are not consistent, however, with at least one large study demonstrating inferiority with respect to therapy with 5-fluorouracil and leucovorin. Raltitrexed has also been used in combination with either oxaliplatin or irinotecan for the treatment of patients with advanced colorectal cancer. These combinations have been associated with acceptable toxicity, response rates in the 35 to 45% range, and overall median survivals of approximately 15 months.385, 386 As has been found to be the case for 5-fluorouracil, intratumoral thymidylate synthase levels have also been demonstrated to predict for responsiveness to raltitrexed as well as overall survival with low levels of thymidylate synthase being associated with higher response rates and longer survival when compared to patients with higher enzyme levels. 387


ZD9331 is a potent quinazoline antifolate inhibitor of TS that does not require polyglutamation for activity. The lack of required polyglutamation of ZD9331, which is in contrast to raltitrexed, may allow for antitumor activity in cells with low FPGS activity. ZD9331 is transported into cells predominantly by the RFC system and competes with both MTX and folinic acid for cellular uptake. Clinical investigations suggest single-agent activity in patients with refractory lung, ovarian and breast cancers.388

Antibody-Directed Enzyme Therapy

Antibody-directed enzyme therapy (ADEPT) systems separate cytotoxic and targeting functions by binding to cell surface markers expressed specifically on malignant cells and activating molecules, including antifolate compounds like MTX at the target cell. This targeted binding and activation theoretically minimizes generalized toxicity secondary to nonspecific delivery of cytotoxic drug. Studies done by Springer et al.389 delivered an antibody-CPG2 enzyme before the nontoxic prodrug CMDA. Once delivered, CMDA was converted to a cytotoxic drug by the action of the localized conjugate at the tumor site. In addition, prodrugs of quinazoline antifolate TS inhibitors (ZD1694 and ICI198583) have been designed and synthesized for use in ADEPT systems. The α-linked L-dipeptide prodrugs were designed to be activated to their corresponding TS inhibitors at the tumor site by prior administration of a monoclonal antibody conjugated to the enzyme carboxypeptidase A. Activation of the α-linked L-alanine dipeptides with carboxypeptidase A led to a cytotoxicity enhancement of 10- to 100-fold.390 ADEPT holds the potential of providing an effective and relatively nontoxic treatment of cancer.391


1. Farber S, Diamond LK, Mercer RD, et al. Temporary remission in acute leukemia in children produced by folic acid antagonist 4-amethopteroylglutamic acid (aminopterin). N Engl J Med 1948;238:787.

2. Hoffmeister RT. Methotrexate therapy in rheumatoid arthritis: 15 years' experience. Am J Med 1983;75:69–73.

3. Rees RB, Bennett JH, Maibach HI, et al. Methotrexate for psoriasis. Arch Dermatol 1967;95:2–11.

4. Calabresi P, Chabner BA. Chemotherapy of neoplastic diseases. In: Gilman AG, Rall TW, Dies DS, et al., eds. The pharmacologic Basis of Therapeutics. 8th Ed. New York: Pergamon Press, 1990:1202.

5. Allegra CJ, Chabner BA, Tuazon CU, et al. Trimetrexate for the treatment of Pneumocystis carinii pneumonia in patients with the acquired immunodeficiency syndrome. N Engl J Med 1987;317:978–985.

6. Allegra CJ, Fine RL, Drake JC, et al. The effect of methotrexate on intracellular folate pools in human MCF-7 breast cancer cells. Evidence for direct inhibition of purine synthesis. J Biol Chem 1986;261:6478–6485.

7. Matherly LH, Barlowe CK, Phillips VM, et al. The effects of 4-amino-antifolates on 5-formyltetrahydrofolate metabolism in L1210 cells. J Biol Chem 1987;262:710–717.

8. Baram J, Allegra CJ, Fine RL, et al. Effect of methotrexate on intracellular folate pools in purified myeloid precursor cells from normal human bone marrow. J Clin Invest 1987;79:692–697.

9. Kesavan V, Sur P, Doig MT, et al. Effects of methotrexate on folates in Krebs ascites and L1210 murine leukemia cells. Cancer Lett 1986;30:55–59.

10. Bunni M, Doig MT, Donato H, et al. Role of methylenetetrahydrofolate depletion in methotrexate-mediated intracellular thymidylate synthesis inhibition in cultured L1210 cells. Cancer Res 1988;48:3398–3404.

11. Seither RL, Trent DF, Mikulecky DC, et al. Folate-pool interconversions and inhibition of biosynthetic processes after exposure of L1210 leukemia cells to antifolates. Experimental and network thermodynamic analyses of the role of dihydrofolate polyglutamylates in antifolate action in cells. J Biol Chem 1989;264:17016–17023.

12. Priest DG, Bunni M, Sirotnak FM. Relationship of reduced folate changes to inhibition of DNA synthesis induced by methotrexate in L1210 cells in vivo. Cancer Res 1989;49:4204–4209.

13. Allegra CJ, Hoang K, Yeh GC, et al. Evidence for direct inhibition of de novo purine synthesis in human MCF-7 breast cells as a principal mode of metabolic inhibition by methotrexate. J Biol Chem 1987;262:13520–13526.

14. Baram J, Chabner BA, Drake JC, et al. Identification and biochemical properties of 10-formyldihydrofolate, a novel folate found in methotrexate-treated cells. J Biol Chem 1988;263: 7105–7111.

15. Kumar P, Kisliuk RL, Gaumont Y, et al. Inhibition of human dihydrofolate reductase by antifolyl polyglutamates. Biochem Pharmacol 1989;38:541–543.

16. Allegra CJ, Chabner BA, Drake JC, et al. Enhanced inhibition of thymidylate synthase by methotrexate polyglutamates. J Biol Chem 1985;260:9720–9726.

17. Allegra CJ, Drake JC, Jolivet J, et al. Inhibition of phosphoribosylaminoimidazolecarboxamide transformylase by methotrexate and dihydrofolic acid polyglutamates. Proc Natl Acad Sci U S A 1985;82:4881–4885.

18. Baggott JE, Vaughn WH, Hudson BB. Inhibition of 5- aminoimidazole-4-carboxamide ribotide transformylase, adenosine deaminase and 5′-adenylate deaminase by polyglutamates of methotrexate and oxidized folates and by 5-aminoimidazole-4-carboxamide riboside and ribotide. Biochem J 1986;236: 193–200.

19. Chu E, Drake JC, Boarman D, et al. Mechanism of thymidylate synthase inhibition by methotrexate in human neoplastic cell lines and normal human myeloid progenitor cells. J Biol Chem 1990;265:8470–8478.

20. Morrison PF, Allegra CJ. Folate cycle kinetics in human breast cancer cells. J Biol Chem 1989;264:10552–10566.

21. Lyons SD, Sant ME, Christopherson RI. Cytotoxic mechanisms of glutamine antagonists in mouse L1210 leukemia. J Biol Chem 1990;265:11377–11381.

22. O'Dwyer PJ, Shoemaker DD, Plowman J, et al. Trimetrexate: a new antifol entering clinical trials. Invest New Drugs 1985;3:71–75.

23. Sigel CW, Macklin AW, Woolley JL Jr, et al. Preclinical biochemical pharmacology and toxicology of piritrexim, a lipophilic inhibitor of dihydrofolate reductase. J Natl Cancer Inst Monogr 1987;5:111–120.

24. Sirotnak FM, DeGraw JI, Schmid FA, et al. New folate analogs of the 10-deaza-aminopterin series. Further evidence for markedly increased antitumor efficacy compared with methotrexate in ascitic and solid murine tumor models. Cancer Chemother Pharmacol 1984;12:26–30.

25. Kamen BA, Eibl B, Cashmore A, et al. Uptake and efficacy of trimetrexate (TMQ, 2,4-diamino-5-methyl-6-[(3,4,5-trimethoxyanilino) methyl] quinazoline), a non-classical antifolate in methotrexate-resistant leukemia cells in vitro. Biochem Pharmacol 1984;33:1697–1699.

26. Jones TR, Calvert AH, Jackman AL, et al. A potent anti- tumor quinazoline inhibitor of thymidylate synthetase, biological properties and therapeutic results in mice. Eur J Cancer 1981;17:11–19.

27. Cheng YC, Dutschman GE, Starnes MC, et al. Activity of the new antifolate N10-propargyl-5,8-dideazafolate and its polyglutamates against human dihydrofolate reductase, human thymidylate synthetase, and KB cells containing different levels of dihydrofolate reductase. Cancer Res 1985;45:598–600.

28. Grindey GB, Shih C, Bernett CJ, et al. A novel pyrrolopyrimidine antifolate that inhibits thymidylate synthase (TS). Am Assoc Cancer Res 1992:2451.

29. Humphreys J, Smith G, Waters K, et al. Antitumor activity of the novel thymidylate synthase inhibitor 1843U89 in cells resistant to antifolates by multiple mechanisms. Am Assoc Cancer Res 1993:1625.

30. Beardsley GP, Taylor EC, Grindley GB, et al. Deaza derivatives of tetrahydrofolic acid: a new class of folate antimetabolite. In: Cooper BA, Whitehead VM, eds. Chemistry and Biology of Pteridines. Berlin: Walter deGruyter, 1986:953.

31. Antony AC, Kane MA, Portillo RM, et al. Studies of the role of a particulate folate-binding protein in the uptake of 5-methyltetrahydrofolate by cultured human KB cells. J Biol Chem 1985; 260:14911–14917.

32. Kamen BA, Capdevila A. Receptor-mediated folate accumulation is regulated by the cellular folate content. Proc Natl Acad Sci US A 1986;83:5983–5987.

33. Fan J, Vitols KS, Huennekens FM. Biotin derivatives of methotrexate and folate. Synthesis and utilization for affinity purification of two membrane-associated folate transporters from L1210 cells. J Biol Chem 1991;266:14862–14865.

34. Brigle KE, Westin EH, Houghton MT, et al. Characterization of two cDNAs encoding folate-binding proteins from L1210 murine leukemia cells. Increased expression associated with a genomic rearrangement. J Biol Chem 1991;266:17243–17249.

35. Chello PL, Sirotnak FM, Dorick DM. Alterations in the kinetics of methotrexate transport during growth of L1210 murine leukemia cells in culture. Mol Pharmacol 1980;18:274–280.

36. Knight CB, Elwood PC, Chabner BA. Future directions for antifolate drug development. Adv Enzyme Regul 1989;29:3–12.

37. Kane MA, Portillo RM, Elwood PC, et al. The influence of extracellular folate concentration on methotrexate uptake by human KB cells. Partial characterization of a membrane- associated methotrexate binding protein. J Biol Chem 1986;261:44–49.

38. Price EM, Freisheim JH. Photoaffinity analogues of methotrexate as folate antagonist binding probes. 2. Transport studies, photoaffinity labeling, and identification of the membrane carrier protein for methotrexate from murine L1210 cells. Biochemistry 1987;26:4757–4763.

39. Henderson GB, Tsuji JM, Kumar HP. Transport of folate compounds by leukemic cells. Evidence for a single influx carrier for methotrexate, 5-methyltetrahydrofolate, and folate in CCRF-CEM human lymphoblasts. Biochem Pharmacol 1987;36:3007–3014.

40. Antony AC. The biological chemistry of folate receptors. Blood 1992;79:2807–2820.

41. Sirotnak FM, Goutas LJ, Jacobsen DM, et al. Carrier-mediated transport of folate compounds in L1210 cells. Initial rate kinetics and extent of duality of entry routes for folic acid and diastereomers of 5-methyltetrahydrohomofolate in the presence of physiological anions. Biochem Pharmacol 1987;36:1659–1667.

42. Matherly LH, Czajkowski CA, Angeles SM. Identification of a highly glycosylated methotrexate membrane carrier in K562 human erythroleukemia cells up-regulated for tetrahydrofolate cofactor and methotrexate transport. Cancer Res 1991;51: 3420–3426.

43. Moscow JA, Gong M, He R, et al. Isolation of a gene encoding a human reduced folate carrier (RFC1) and analysis of its expression in transport-deficient, methotrexate-resistant human breast cancer cells. Cancer Res 1995;55:3790–3794.

44. Prasad PD, Ramamoorthy S, Leibach FH, et al. Molecular cloning of the human placental folate transporter. Biochem Biophys Res Commun 1995;206:681–687.

45. Zhao R, Gao F, Wang PJ etal. Role of the amino acid 45 residue in reduced folate carrier function and ion-dependent transport as characterized by site-directed mutagenesis. Mol Pharmacol 2000;57(2):317–323.

46. Zhao R, Wang PJ, Gao F, etal. Residues 45 and 404 in the murine reduced folate carrier may interact to alter carrier binding and mobility. Biochim Biophys Acta 2003;1613(1–2):49–56.

47. Sharina IG, Zhao R, Wang Y, et al. Mutational analysis of the functional role of conserved arginine and lysine residues in transmembrane domains of the murine reduced folate carrier. Mol Pharmacol 2001;59(5):1022–1028.

48. Zhao R, Gao F, Hanscom M, et al. A prominent low-pH methotrexate transport activity in human solid tumors: contribution to the preservation of methotrexate pharmacologic activity in HeLa cells lacking the reduced folate carrier. Clin Cancer Res 2004;10(2):718–727.

49. Campbell IG, Jones TA, Foulkes WD, et al. Folate-binding protein is a marker for ovarian cancer. Cancer Res 1991;51:5329–5338.

50. Coney LR, Tomassetti A, Carayannopoulos L, et al. Cloning of a tumor-associated antigen: MOv18 and MOv19 antibodies recognize a folate-binding protein. Cancer Res 1991;51:6125–6132.

51. Elwood PC, Kane MA, Portillo RM, et al. The isolation, characterization, and comparison of the membrane-associated and soluble folate-binding proteins from human KB cells. J Biol Chem 1986;261:15416–15423.

52. Shen F, Ross JF, Wang X, et al. Identification of a novel folate receptor, a truncated receptor, and receptor type beta in hematopoietic cells: cDNA cloning, expression, immunoreactivity, and tissue specificity. Biochemistry 1994;33:1209–1215.

53. Roberts SJ, Petropavlovskaja M, Chung KN, et al. Role of individual N-linked glycosylation sites in the function and intracellular transport of the human alpha folate receptor. Arch Biochem Biophys 1998;351:227–235.

54. Elwood PC, Nachmanoff K, Saikawa Y, et al. The divergent 5′ termini of the alpha human folate receptor (hFR) mRNAs originate from two tissue-specific promoters and alternative splicing: characterization of the alpha hFR gene structure. Biochemistry 1997;36:1467–1478.

55. Roberts SJ, Chung KN, Nachmanoff K, et al. Tissue-specific promoters of the alpha human folate receptor gene yield transcripts with divergent 5′ leader sequences and different translational efficiencies. Biochem J 1997;326:439–447.

56. Sun XL, Murphy BR, Li QJ, et al. Transduction of folate receptor cDNA into cervical carcinoma cells using recombinant adeno-associated virions delays cell proliferation in vitro and in vivo. J Clin Invest 1995;96:1535–1547.

57. Shen F, Zheng X, Wang J, et al. Identification of amino acid residues that determine the differential ligand specificities of folate receptors alpha and beta. Biochemistry 1997;36:6157–6163.

58. Wu M, Fan J, Gunning W, et al. Clustering of GPI-anchored folate receptor independent of both cross-linking and association with caveolin. J Membr Biol 1997;159:137–147.

59. Anderson RG, Kamen BA, Rothberg KG, et al. Potocytosis: sequestration and transport of small molecules by caveolae. Science 1992;255:410–411.

60. Kamen BA, Smith AK, Anderson RG. The folate receptor works in tandem with a probenecid-sensitive carrier in MA104 cells in vitro. J Clin Invest 1991;87:1442–1449.

61. Chang WJ, Rothberg KG, Kamen BA, et al. Lowering the cholesterol content of MA104 cells inhibits receptor-mediated transport of folate. J Cell Biol 1992;118:63–69.

62. Smart EJ, Mineo C, Anderson RG. Clustered folate receptors deliver 5-methyltetrahydrofolate to cytoplasm of MA104 cells. J Cell Biol 1996;134:1169–1177.

63. Spinella MJ, Brigle KE, Sierra EE, et al. Distinguishing between folate receptor-alpha-mediated transport and reduced folate carrier-mediated transport in L1210 leukemia cells. J Biol Chem 1995;270:7842–7849.

64. Westerhof GR, Rijnboutt S, Schornagel JH, et al. Functional activity of the reduced folate carrier in KB, MA104, and IGROV-I cells expressing folate-binding protein. Cancer Res 1995;55: 3795–3802.

65. Schuetz JD, Matherly LH, Westin EH, et al. Evidence for a functional defect in the translocation of the methotrexate transport carrier in a methotrexate-resistant murine L1210 leukemia cell line. J Biol Chem 1988;263:9840–9847.

66. Assaraf YG, Schimke RT. Identification of methotrexate transport deficiency in mammalian cells using fluoresceinated methotrexate and flow cytometry. Proc Natl Acad Sci U S A 1987;84:7154–7158.

67. Rodenhuis S, McGuire JJ, Narayanan R, et al. Development of an assay system for the detection and classification of methotrexate resistance in fresh human leukemic cells. Cancer Res 1986;46:6513–6519.

68. Schuetz JD, Westin EH, Matherly LH, et al. Membrane protein changes in an L1210 leukemia cell line with a translocation defect in the methotrexate-tetrahydrofolate cofactor transport carrier. J Biol Chem 1989;264:16261–16267.

69. Jansen G, Westerhof GR, Kathmann I, et al. Identification of a membrane-associated folate-binding protein in human leukemic CCRF-CEM cells with transport-related methotrexate resistance [published erratum appears in Cancer Res 1995;55 (18):4203]. Cancer Res 1989;49:2455–2459.

70. Zhao R, Assaraf YG, Goldman ID. A mutated murine reduced folate carrier (RFC1) with increased affinity for folic acid, decreased affinity for methotrexate, and an obligatory anion requirement for transport function. J Biol Chem 1998;273: 19065–19071.

71. Zhao R, Assaraf YG, Goldman ID. A reduced folate carrier mutation produces substrate-dependent alterations in carrier mobility in murine leukemia cells and methotrexate resistance with conservation of growth in 5-formyltetrahydrofolate. J Biol Chem 1998;273:7873–7879.

72. Gifford AJ, Haber M, Witt TL etal. Role of the E45K-reduced folate carrier gene mutation in methotrexate resistance in human leukemia cells. Leukemia 2002;16(12):2379–2387.

73. Trippett T, Schlemmer S, Elisseyeff Y, et al. Defective transport as a mechanism of acquired resistance to methotrexate in patients with acute lymphoblastic leukemia. Blood 1992;80:1158–1162.

74. Levy AS, Sather HN, Steinherz PG etal. Reduced folate carrier and dihydrofolate reductase expression in acucte lymphocytic leukemia may predict outcome: a Children's Cancer Group Study. J Pediatr Hematol Oncol 2003;25(9):688–695.

75. Guo W, Healey JH, Meyers PA, et al. Mechanisms of methotrexate resistance in osteosarcoma. Clin Cancer Res 1999;5: 621–627.

76. Ifergan I, Meller I, Issakov J etal. Reduced folate carrier protein expression in osteosarcoma: implications for the prediction of tumor chemosensitivity. Cancer 2003;98(9):1958–1966.

77. Liu S, Song L, Bevins R, et al. The murine-reduced folate carrier gene can act as a selectable marker and a suicide gene in hematopoietic cells in vivo. Hum Gene Ther 2002;13(14): 1777–1782.

78. Moscow JA. Methotrexate transport and resistance. Leuk Lymphoma 1998;30:215–224.

79. Matherly LH, Goldman DI. Membrane transport of folates. Vitam Horm 2003;66:403–456.

80. Taylor IW, Slowiaczek P, Friedlander ML, et al. Selective toxicity of a new lipophilic antifolate, BW301U, for methotrexate-resistant cells with reduced drug uptake. Cancer Res 1985;45:978–982.

81. Mini E, Moroson BA, Franco CT, et al. Cytotoxic effects of folate antagonists against methotrexate-resistant human leukemic lymphoblast CCRF-CEM cell lines. Cancer Res 1985;45:325–330.

82. Schmid FA, Sirotnak FM, Otter GM, et al. Combination chemotherapy with a new folate analog: activity of 10- ethyl-10-deaza-aminopterin compared to methotrexate with 5-fluorouracil and alkylating agents against advanced metastatic disease in murine tumor models. Cancer Treat Rep 1987;71: 727–732.

83. Jansen G, Schornagel JH, Westerhof GR, et al. Multiple membrane transport systems for the uptake of folate-based thymidylate synthase inhibitors. Cancer Res 1990;50:7544–7548.

84. Westerhof GR, Jansen G, van Emmerik N, et al. Membrane transport of natural folates and antifolate compounds in murine L1210 leukemia cells: role of carrier- and receptor- mediated transport systems. Cancer Res 1991;51:5507–5513.

85. Wang Y, Zhao R, Goldman ID. Decreased expression of the reduced folate carrier and folylpolyglutamate synthetase is the basis for acquired resistance to the pemetrexed antifolate (LY231514) in an L1210 murine leukemia cell line. Biochem Pharmacol 2003;65(7):1163–1170.

86. Zhao R, Hanscom M, Chattopadhyay S, et al. Selective Preservation of pemetrexed pharmacological activity in HeLa cells lacking the reduced folate carrier: association with the presence of a secondary transport pathway. Cancer Res 2004;64(9):3313–3319.

87. Henderson GB, Strauss BP. Growth inhibition by homofolate in tumor cells utilizing a high-affinity folate binding protein as a means for folate internalization. Biochem Pharmacol 1990;39:2019–2025.

88. Mendelsohn LG, Gates SB, Habeck LL, et al. The role of dietary folate in modulation of folate receptor expression, folylpolyglutamate synthetase activity and the efficacy and toxicity of lometrexol. Adv Enzyme Regul 1996;36:365–381.

89. Lin JT, Cashmore AR, Baker M, et al. Phase I studies with trimetrexate: clinical pharmacology, analytical methodology, and pharmacokinetics. Cancer Res 1987;47:609–616.

90. Laszlo J, Brenckman WD Jr, Morgan E, et al. Initial clinical studies of piritrexim. J Natl Cancer Inst Monogr 1987;5:121–125.

91. Allegra CJ, Kovacs JA, Drake JC, et al. Activity of antifolates against Pneumocystis carinii dihydrofolate reductase and identification of a potent new agent. J Exp Med 1987;165:926–931.

92. Currie VE, Warrell RP Jr, Arlin Z, et al. Phase I trial of 10- deaza-aminopterin in patients with advanced cancer. Cancer Treat Rep 1983;67:149–154.

93. Casper ES, Christman KL, Schwartz GK, et al. Edatrexate in patients with soft tissue sarcoma. Activity in malignant fibrous histiocytoma. Cancer 1993;72:766–770.

94. Vandenberg TA, Pritchard KI, Eisenhauer EA, et al. Phase II study of weekly edatrexate as first-line chemotherapy for metastatic breast cancer: a National Cancer Institute of Canada Clinical Trials Group study. J Clin Oncol 1993;11:1241–1244.

95. Kuriakose P, Gandara DR, and Perez EA. Phase I trial of edatrexate in advanced breast and other cancers. Cancer Invest 2002;20(4):473–749.

96. Henderson GB, Zevely EM. Inhibitory effects of probenecid on the individual transport routes which mediate the influx and efflux of methotrexate in L1210 cells. Biochem Pharmacol 1985;34:1725–1729.

97. Henderson GB, Tsuji JM. Methotrexate efflux in L1210 cells. Kinetic and specificity properties of the efflux system sensitive to bromosulfophthalein and its possible identity with a system which mediates the efflux of 3′,5′-cyclic AMP. J Biol Chem 1987;262:13571–13578.

98. Henderson GB, Tsuji JM, Kumar HP. Characterization of the individual transport routes that mediate the influx and efflux of methotrexate in CCRF-CEM human lymphoblastic cells. Cancer Res 1986;46:1633–1638.

99. Zeng H, Liu G, Rea PA, et al. Transport of amphipathic anions by human multidrug resistance protein 3. Cancer Res 2000;60(17):4779–4784.

100. Zeng H, Chen ZS, Belinsky MG, et al. Transport of methotrexate (MTX) and folates by multidrug resistance protein (MRP) 3 and MRP1: effect of polyglutamation on MTX transport. Cancer Res 2001;61(19):7225–7232.

101. Chen ZS, Lee K, Walther S, et al. Analysis of methotrexate and folate transport by multidrug resistance protein 4 (ABCC4): MRP4 is a component of the methotrexate efflux system. Cancer Res 2002;62(11):3144–3150.

102. Assaraf YG, Rothem L, Hooijberg JH, et al. Loss of multidrug resistance protein 1 expression and folate efflux activity results i a highly concentrative folate transport in human leukemia cells. J Biol Chem 2003;278(9):6680–6686.

103. Sirotnak FM, Wendel HG, Bornmann WG, et al. Co-administration of probenecid, an inhibitor of a cMOAT/MRP-like plasma membrane ATPase, greatly enhanced the efficacy of a new 10-deazaaminopterin against human solid tumors in vivo. Clin Cancer Res 2000;6(9):3705–3712.

104. Stark M, Rothem L, Jansen G, et al. Antifolate resistance associated with loss of MRP1 expression and function in Chinese hamster ovary cells with markedly impaired export of folate and cholate. Mol Pharmacol 2003;64(2):220–227.

105. Volk EL, Farley KM, Wu Y, et al. Overexpression of wild-type breast cancer resistance protein mediates methotrexate resistance. Cancer Res 2002;62(17):5035–5040.

106. Volk EL, Schneider E. Wild-type breast cancer resistance protein (BCRP/ABCG2) is a methotrexate polyglutamate transporter. Cancer Res 2003;63(17):5538–5543.

107. Chen ZS, Robey RW, Belinsky MG, et al. Transpor of methotrexate polyglutamates, and 17beta-estradiol 17-(beta-D-glucuronide) by ABCG2: effects of acquired mutations as R482 on methotrexate transport. Cancer Res 2003;63(14):4048–4054.

108. Koizumi S, Curt GA, Fine RL, et al. Formation of methotrexate polyglutamates in purified myeloid precursor cells from normal human bone marrow. J Clin Invest 1985;75:1008–1014.

109. Fabre I, Fabre G, Goldman ID. Polyglutamylation, an important element in methotrexate cytotoxicity and selectivity in tumor versus murine granulocytic progenitor cells in vitro. Cancer Res 1984;44:3190–3195.

110. Schilsky RL, Bailey BD, Chabner BA. Methotrexate polyglutamate synthesis by cultured human breast cancer cells. Proc Natl Acad Sci U S A 1980;77:2919–2922.

111. Jolivet J, Schilsky RL, Bailey BD, et al. Synthesis, retention, and biological activity of methotrexate polyglutamates in cultured human breast cancer cells. J Clin Invest 1982;70:351–360.

112. Kennedy DG, Van den Berg HW, Clarke R, et al. The effect of the rate of cell proliferation on the synthesis of methotrexate poly-gamma-glutamates in two human breast cancer cell lines. Biochem Pharmacol 1985;34:3087–3090.

113. Jolivet J, Chabner BA. Intracellular pharmacokinetics of methotrexate polyglutamates in human breast cancer cells. Selective retention and less dissociable binding of 4-NH2- 10-CH3-pteroylglutamate4 and 4-NH2-10-CH3-pteroylglutamate5 to dihydrofolate reductase. J Clin Invest 1983;72:773–778.

114. Winick NJ, Kamen BA, Balis FM, et al. Folate and methotrexate polyglutamate tissue levels in rhesus monkeys following chronic low-dose methotrexate. Cancer Drug Deliv 1987;4:25–31.

115. Shane B. Folylpolyglutamate synthesis and role in the regulation of one-carbon metabolism. Vitam Horm 1989;45:263–335.

116. Gewirtz DA, White JC, Randolph JK, et al. Formation of methotrexate polyglutamates in rat hepatocytes. Cancer Res 1979;39:2914–2918.

117. Clarke L, Waxman DJ. Human liver folylpolyglutamate synthetase: biochemical characterization and interactions with folates and folate antagonists. Arch Biochem Biophys 1987;256:585–596.

118. Cichowicz DJ, Shane B. Mammalian folylpoly-gamma- glutamate synthetase. 1. Purification and general properties of the hog liver enzyme. Biochemistry 1987;26:504–512.

119. Cichowicz DJ, Shane B. Mammalian folylpoly-gamma- glutamate synthetase. 2. Substrate specificity and kinetic properties. Biochemistry 1987;26:513–521.

120. McGuire JJ, Hsieh P, Franco CT, et al. Folylpolyglutamate synthetase inhibition and cytotoxic effects of methotrexate analogs containing 2, omega-diaminoalkanoic acids. Biochem Pharmacol 1986;35:2607–2613.

121. Galivan J, Johnson T, Rhee M, et al. The role of folylpolyglutamate synthetase and gamma-glutamyl hydrolase in altering cellular folyl- and antifolylpolyglutamates. Adv Enzyme Regul 1987;26:147–155.

122. Panetta JC, Wall A, Pui CH, et al. Methotrexate intracellular disposition in acute lymphoblastic leukemia: a mathematical model of gamma-glutamyl hydrolase activity. Clin Cancer Res 2002;8(7):2423–2429.

123. Yao R, Schneider E, Ryan TJ, et al. Human gamma- glutamyl hydrolase: cloning and characterization of the enzyme expressed in vitro. Proc Natl Acad Sci U S A 1996;93:10134–10138.

124. Cole PD, Kamen BA, Gorlick R, et al. Effects of overexpression of gamma-glutamyl hydrolase on methotrexate metabolism and resistance. Cancer Res 2001;61(11):4599–4604.

125. Longo GS, Gorlick R, Tong WP, et al. Disparate affinities of antifolates for folylpolyglutamate synthetase from human leukemia cells. Blood 1997;90:1241–1245.

126. Sirotnak FM, Chello PL, Piper JR, et al. Growth inhibitory, transport and biochemical properties of the gamma- glutamyl and gamma-aspartyl peptides of methotrexate in L1210 leukemia cells in vitro. Biochem Pharmacol 1978;27:1821–1825.

127. Galivan J, Nimec Z, Balinska M. Regulation of methotrexate polyglutamate accumulation in vitro: effects of cellular folate content. Biochem Pharmacol 1983;32:3244–3247.

128. Jolivet J, Faucher F, Pinard MF. Influence of intracellular folates on methotrexate metabolism and cytotoxicity. Biochem Pharmacol 1987;36:3310–3312.

129. Jolivet J, Cole DE, Holcenberg JS, et al. Prevention of methotrexate cytotoxicity by asparaginase inhibition of methotrexate polyglutamate formation. Cancer Res 1985;45:217–220.

130. Sur P, Fernandes DJ, Kute TE, et al. L-asparaginase-induced modulation of methotrexate polyglutamylation in murine leukemia L5178Y. Cancer Res 1987;47:1313–1318.

131. Matherly LH, Fry DW, Goldman ID. Role of methotrexate polyglutamylation and cellular energy metabolism in inhibition of methotrexate binding to dihydrofolate reductase by 5-formyltetrahydrofolate in Ehrlich ascites tumor cells in vitro. Cancer Res 1983;43:2694–2699.

132. Allegra CJ, Drake JC, Jolivet J, et al. Inhibition of folate-dependent enzymes by methotrexate polyglutamates. In: Goldman ID, ed. Proceedings of the Second Workshop on Folyl and Antifolyl Polyglutamates. New York: Praeger, 1985:348–359.

133. Galivan J, Inglese J, McGuire JJ, et al. Gamma-fluoromethotrexate: synthesis and biological activity of a potent inhibitor of dihydrofolate reductase with greatly diminished ability to form poly-gamma-glutamates. Proc Natl Acad Sci U S A 1985;82: 2598–2602.

134. Samuels LL, Moccio DM, Sirotnak FM. Similar differential for total polyglutamylation and cytotoxicity among various folate analogues in human and murine tumor cells in vitro. Cancer Res 1985;45:1488–1495.

135. Matherly LH, Voss MK, Anderson LA, et al. Enhanced polyglutamylation of aminopterin relative to methotrexate in the Ehrlich ascites tumor cell in vitro. Cancer Res 1985;45:1073–1078.

136. Cowan KH, Jolivet J. A methotrexate-resistant human breast cancer cell line with multiple defects, including diminished formation of methotrexate polyglutamates. J Biol Chem 1984;259:10793–10800.

137. Mauritz R, Peters GJ, Priest DG, et al. Multiple mechanisms of resistance to methotrexate and novel antifolates in human CCRF-CEM leukemia cells and their implications for folate homeostasis. Biochem Pharmacol 2002;63(2):105–115.

138. Liani E, Rothem L, Bunni MA, et al. Loss of folylpoly-gamma-glutamate synthetase activity is a dominant mechanism of resistance to polyglutamation-dependent novel antifolates in multiple human leukemia sublines. Int J Cancer 2003;103(5): 587–599.

139. Pizzorno G, Mini E, Coronnello M, et al. Impaired polyglutamylation of methotrexate as a cause of resistance in CCRF-CEM cells after short-term, high-dose treatment with this drug. Cancer Res 1988;48:2149–2155.

140. Pizzorno G, Chang YM, McGuire JJ, et al. Inherent resistance of human squamous carcinoma cell lines to methotrexate as a result of decreased polyglutamylation of this drug. Cancer Res 1989;49:5275–5280.

141. McCloskey DE, McGuire JJ, Russell CA, et al. Decreased folylpolyglutamate synthetase activity as a mechanism of methotrexate resistance in CCRF-CEM human leukemia sublines. J Biol Chem 1991;266:6181–6187.

142. Faessel HM, Slocum HK, Jackson RC, et al. Super in vitro synergy between inhibitors of dihydrofolate reductase and inhibitors of other folate-requiring enzymes: the critical role of polyglutamylation. Cancer Res 1998;58:3036–3050.

143. Pizzorno G, Sokoloski JA, Cashmore AR, et al. Intracellular metabolism of 5,10-dideazatetrahydrofolic acid in human leukemia cell lines. Mol Pharmacol 1991;39:85–89.

144. Curt GA, Jolivet J, Carney DN, et al. Determinants of the sensitivity of human small-cell lung cancer cell lines to methotrexate. J Clin Invest 1985;76:1323–1329.

145. Longo GS, Gorlick R, Tong WP, et al. Gamma-glutamyl hydrolase and folylpolyglutamate synthetase activities predict polyglutamylation of methotrexate in acute leukemias. Oncol Res 1997;9:259–263.

146. Galpin AJ, Schuetz JD, Masson E, et al. Differences in folylpolyglutamate synthetase and dihydrofolate reductase expression in human B-lineage versus T-lineage leukemic lymphoblasts: mechanisms for lineage differences in methotrexate polyglutamylation and cytotoxicity. Mol Pharmacol 1997;52:155–163.

147. Panetta JC, Yanishevski Y, Pui CH, et al. A mathematical model of in vivo methotrexate accumulation in acute lymphoblastic leukemia. Cancer Chemother Pharmacol 2002;50(5):419–428.

148. Mantadakis E, Smith AK, Hynan L etal. Methotrexate polyglutamation may lack prognostic significance in children with B-cell precursor acute lymphoblastic leukemia treated with intensive oral methotrexate. J Pediatr Hematol Oncol 2002;24(8): 736–642.

149. Li WW, Lin JT, Tong WP, et al. Mechanisms of natural resistance to antifolates in human soft tissue sarcomas. Cancer Res 1992;52:1434–1438.

150. Li WW, Lin JT, Schweitzer BI, et al. Intrinsic resistance to methotrexate in human soft tissue sarcoma cell lines. Cancer Res 1992;52:3908–3913.

151. Matthews DA, Alden RA, Bolin JT, et al. X-ray structural studies of dihydrofolate reductase. In: Kisliuk RL, Brown GM, eds. Chemistry and Biology of Pteridines. New York: Elsevier/North Holland, 1979:465.

152. Matthews DA, Alden RA, Bolin JT, et al. Dihydrofolate reductase: x-ray structure of the binary complex with methotrexate. Science 1977;197:452–455.

153. Appleman JR, Howell EE, Kraut J, et al. Role of aspartate 27 in the binding of methotrexate to dihydrofolate reductase from Escherichia coli. J Biol Chem 1988;263:9187–9198.

154. Taira K, Benkovic SJ. Evaluation of the importance of hydrophobic interactions in drug binding to dihydrofolate reductase. J Med Chem 1988;31:129–137.

155. Oefner C, D'Arcy A, Winkler FK. Crystal structure of human dihydrofolate reductase complexed with folate. Eur J Biochem 1988;174:377–385.

156. Cody V, Ciszak E. Computer graphic modeling in drug design—conformational analysis of antifolate binding to avian dihydrofolate reductase: crystal and molecular structures of 2,4-diamino-5-cyclohexyl-6-methylpyrimidine and 5-cyclohexyl-6-methyluracil. Anticancer Drug Des 1991;6:83–93.

157. Freisheim JH, Kumar AA, Blankenship D. Structure-function relationships of dihydrofolate reductases: sequence homology considerations and active center residues. In: Kislink RL, Brown GM, eds. Chemistry and Biology of Pteridines. New York: Elsevier/North Holland, 1979:419.

158. Schweitzer BI, Srimatkandada S, Gritsman H, et al. Probing the role of two hydrophobic active site residues in the human dihydrofolate reductase by site-directed mutagenesis. J Biol Chem 1989;264:20786–20795.

159. Thompson PD, Freisheim JH. Conversion of arginine to lysine at position 70 of human dihydrofolate reductase: generation of a methotrexate-insensitive mutant enzyme. Biochemistry 1991;30:8124–8130.

160. Dicker AP, Waltham MC, Volkenandt M, et al. Methotrexate resistance in an in vivo mouse tumor due to a non- active-site dihydrofolate reductase mutation. Proc Natl Acad Sci U S A 1993;90:11797–11801.

161. Bystroff C, Kraut J. Crystal structure of unliganded Escherichia coli dihydrofolate reductase. Ligand-induced conformational changes and cooperativity in binding. Biochemistry 1991;30:2227–2239.

162. Zhao SC, Banerjee D, Mineishi S, et al. Post-transplant methotrexate administration leads to improved curability of mice bearing a mammary tumor transplanted with marrow transduced with a mutant human dihydrofolate reductase cDNA. Hum Gene Ther 1997;8:903–909.

163. Flasshove M, Banerjee D, Leonard JP, et al. Retroviral transduction of human CD34+ umbilical cord blood progenitor cells with a mutated dihydrofolate reductase cDNA. Hum Gene Ther 1998;9:63–71.

164. Mareya SM, Sorrentino BP, Blakley RL. Protection of CCRF-CEM human lymphoid cells from antifolates by retroviral gene transfer of variants of murine dihydrofolate reductase. Cancer Gene Ther 1998;5:225–235.

165. Appleman JR, Prendergast N, Delcamp TJ, et al. Kinetics of the formation and isomerization of methotrexate complexes of recombinant human dihydrofolate reductase. J Biol Chem 1988;263:10304–10313.

166. Kamen BA, Whyte-Bauer W, Bertino JR. A mechanism of resistance to methotrexate. NADPH but not NADH stimulation of methotrexate binding to dihydrofolate reductase. Biochem Pharmacol 1983;32:1837–1841.

167. Jackson RC, Hart LI, Harrap KR. Intrinsic resistance to methotrexate of cultured mammalian cells in relation to the inhibition kinetics of their dihydrofolate reductases. Cancer Res 1976;36:1991–1997.

168. Kumar P, Kisliuk RL, Gaumont Y, et al. Interaction of polyglutamyl derivatives of methotrexate, 10-deazaaminopterin, and dihydrofolate with dihydrofolate reductase. Cancer Res 1986;46:5020–5023.

169. Blakley RL, Cocco L. Role of isomerization of initial complexes in the binding of inhibitors to dihydrofolate reductase. Biochemistry 1985;24:4772–4777.

170. White JC. Reversal of methotrexate binding to dihydrofolate reductase by dihydrofolate. Studies with pure enzyme and computer modeling using network thermodynamics. J Biol Chem 1979;254:10889–10895.

171. Allegra CJ, Boarman D. Interaction of methotrexate polyglutamates and dihydrofolate during leucovorin rescue in a human breast cancer cell line (MCF-7). Cancer Res 1990; 50:3574–3578.

172. Cohen M, Bender RA, Donehower R, et al. Reversibility of high-affinity binding of methotrexate in L1210 murine leukemia cells. Cancer Res 1978;38:2866–2870.

173. White JC, Loftfield S, Goldman ID. The mechanism of action of methotrexate. III. Requirement of free intracellular methotrexate for maximal suppression of (14C)formate incorporation into nucleic acids and protein. Mol Pharmacol 1975;11:287–297.

174. Galivan J. Evidence for the cytotoxic activity of polyglutamate derivatives of methotrexate. Mol Pharmacol 1980;17:105–110.

175. Drake JC, Allegra CJ, Baram J, et al. Effects on dihydrofolate reductase of methotrexate metabolites and intracellular folates formed following methotrexate exposure of human breast cancer cells. Biochem Pharmacol 1987;36:2416–2418.

176. Boarman DM, Baram J, Allegra CJ. Mechanism of leucovorin reversal of methotrexate cytotoxicity in human MCF-7 breast cancer cells. Biochem Pharmacol 1990;40:2651–2660.

177. Kruger-McDermott C, Balinska M, Galivan J. Dihydrofolate-mediated reversal of methotrexate toxicity to hepatoma cells in vitro. Cancer Lett 1986;30:79–84.

178. Goldie JH, Dedhar S, Krystal G. Properties of a methotrexate-insensitive variant of dihydrofolate reductase derived from methotrexate-resistant L5178Y cells. J Biol Chem 1981;256: 11629–11635.

179. McIvor RS, Simonsen CC. Isolation and characterization of a variant dihydrofolate reductase cDNA from methotrexate- resistant murine L5178Y cells. Nucleic Acids Res 1990;18:7025–7032.

180. Flintoff WF, Essani K. Methotrexate-resistant Chinese hamster ovary cells contain a dihydrofolate reductase with an altered affinity for methotrexate. Biochemistry 1980;19:4321–4327.

181. Melera PW, Davide JP, Hession CA, et al. Phenotypic expression in Escherichia coli and nucleotide sequence of two Chinese hamster lung cell cDNAs encoding different dihydrofolate reductases. Mol Cell Biol 1984;4:38–48.

182. Melera PW, Lewis JA, Biedler JL, et al. Antifolate-resistant Chinese hamster cells. Evidence for dihydrofolate reductase gene amplification among independently derived sublines overproducing different dihydrofolate reductases. J Biol Chem 1980;255:7024–7028.

183. Haber DA, Schimke RT. Unstable amplification of an altered dihydrofolate reductase gene associated with double- minute chromosomes. Cell 1981;26:355–362.

184. Cowan KH, Goldsmith ME, Levine RM, et al. Dihydrofolate reductase gene amplification and possible rearrangement in estrogen-responsive methotrexate-resistant human breast cancer cells. J Biol Chem 1982;257:15079–15086.

185. Melera PW, Davide JP, Oen H. Antifolate-resistant Chinese hamster cells. Molecular basis for the biochemical and structural heterogeneity among dihydrofolate reductases produced by drug-sensitive and drug-resistant cell lines. J Biol Chem 1988;263:1978–1990.

186. Dedhar S, Hartley D, Fitz-Gibbons D, et al. Heterogeneity in the specific activity and methotrexate sensitivity of dihydrofolate reductase from blast cells of acute myelogenous leukemia patients. J Clin Oncol 1985;3:1545–1552.

187. Takebe N, Nakahara S, Zhao SC, et al. Comparison of methotrexate resistance conferred by a mutated dihydrofolate reductase cDNA in two different retroviral vectors. Cancer Gene Ther 2000;7(6):910–919.

188. Meisel R, Bardenheuer W, Strehblow C, et al. Efficient protection from methotrexate toxicity and selection of transduced human hematopoietic cells following gene transfer of dihydrofolate reductase mutants. Exp Hematol 2003;31(12):1215–1222.

189. Hamlin JL, Biedler JL. Replication pattern of a large homogenously staining chromosome region in antifolate-resistant Chinese hamster cell lines. J Cell Physiol 1981;107:101–114.

190. Brown PC, Beverley SM, Schimke RT. Relationship of amplified dihydrofolate reductase genes to double minute chromosomes in unstably resistant mouse fibroblast cell lines. Mol Cell Biol 1981;1:1077–1083.

191. Meltzer PS, Cheng YC, Trent JM. Analysis of dihydrofolate reductase gene amplification in a methotrexate-resistant human tumor cell line. Cancer Genet Cytogenet 1985;17:289–300.

192. Singer MJ, Mesner LD, Friedman CL, et al. Amplification of the human dihydrofolate reductase gene via double minutes is initiated by chromosome breaks. Proc Natl Acad Sci USA 2000;97(14):7921–7926.

193. Hoy CA, Rice GC, Kovacs M, et al. Over-replication of DNA in S phase Chinese hamster ovary cells after DNA synthesis inhibition. J Biol Chem 1987;262:11927–11934.

194. Fanin R, Banerjee D, Volkenandt M, et al. Mutations leading to antifolate resistance in Chinese hamster ovary cells after exposure to the alkylating agent ethylmethanesulfonate. Mol Pharmacol 1993;44:13–21.

195. Sharma RC, Schimke RT. Enhancement of the frequency of methotrexate resistance by gamma-radiation in Chinese hamster ovary and mouse 3T6 cells. Cancer Res 1989;49:3861–3866.

196. Barsoum J, Varshavsky A. Mitogenic hormones and tumor promoters greatly increase the incidence of colony-forming cells bearing amplified dihydrofolate reductase genes. Proc Natl Acad Sci U S A 1983;80:5330–5334.

197. Newman EM, Lu Y, Kashani-Sabet M, et al. Mechanisms of cross-resistance to methotrexate and 5-fluorouracil in an A2780 human ovarian carcinoma cell subline resistant to cisplatin. Biochem Pharmacol 1988;37:443–447.

198. Rice GC, Ling V, Schimke RT. Frequencies of independent and simultaneous selection of Chinese hamster cells for methotrexate and doxorubicin (Adriamycin) resistance. Proc Natl Acad Sci U S A 1987;84:9261–9264.

199. Schuetz JD, Gorse KM, Goldman ID, et al. Transient inhibition of DNA synthesis by 5-fluorodeoxyuridine leads to overexpression of dihydrofolate reductase with increased frequency of methotrexate resistance. J Biol Chem 1988;263:7708–7712.

200. Wright JA, Smith HS, Watt FM, et al. DNA amplification is rare in normal human cells. Proc Natl Acad Sci U S A 1990;87:1791–1795.

201. Sowers R, Toguchida J, Qin J etal. MRNA expression levels of E2F transcription factors correlate with dihydrofolate reductase, reduced folate carrier, and thymidylate synthase mRNA expression in osteosarcoma. Mol Cancer Ther 2003;2(6):535–541.

202. Cowan KH, Goldsmith ME, Ricciardone MD, et al. Regulation of dihydrofolate reductase in human breast cancer cells and in mutant hamster cells transfected with a human dihydrofolate reductase minigene. Mol Pharmacol 1986;30:69–76.

203. Chu E, Takimoto CH, Voeller D, et al. Specific binding of human dihydrofolate reductase protein to dihydrofolate reductase messenger RNA in vitro. Biochemistry 1993;32:4756–4760.

204. Ercikan-Abali EA, Banerjee D, Waltham MC, et al. Dihydrofolate reductase protein inhibits its own translation by binding to dihydrofolate reductase mRNA sequences within the coding region. Biochemistry 1997;36:12317–12322.

205. Curt GA, Carney DN, Cowan KH, et al. Unstable methotrexate resistance in human small-cell carcinoma associated with double minute chromosomes. N Engl J Med 1983;308:199–202.

206. Curt GA, Jolivet J, Bailey BD, et al. Synthesis and retention of methotrexate polyglutamates by human small cell lung cancer. Biochem Pharmacol 1984;33:1682–1685.

207. Matherly LH, Taub JW, Wong SC, et al. Increased frequency of expression of elevated dihydrofolate reductase in T-cell versus B-precursor acute lymphoblastic leukemia in children. Blood 1997;90:578–589.

208. Seither RL, Rape TJ, Goldman ID. Further studies on the pharmacologic effects of the 7-hydroxy catabolite of methotrexate in the L1210 murine leukemia cell. Biochem Pharmacol 1989;38:815–822.

209. Rhee MS, Balinska M, Bunni M, et al. Role of substrate depletion in the inhibition of thymidylate biosynthesis by the dihydrofolate reductase inhibitor trimetrexate in cultured hepatoma cells. Cancer Res 1990;50:3979–3984.

210. Rhee MS, Coward JK, Galivan J. Depletion of 5,10-methylenetetrahydrofolate and 10-formyltetrahydrofolate by methotrexate in cultured hepatoma cells. Mol Pharmacol 1992;42:909–916.

211. Trent DF, Seither RL, Goldman ID. Compartmentation of intracellular folates. Failure to interconvert tetrahydrofolate cofactors to dihydrofolate in mitochondria of L1210 leukemia cells treated with trimetrexate [published erratum appears in Biochem Pharmacol 1991;42(12):2405]. Biochem Pharmacol 1991;42:1015–1019.

212. Li JC, Kaminskas E. Accumulation of DNA strand breaks and methotrexate cytotoxicity. Proc Natl Acad Sci U S A 1984;81: 5694–5698.

213. Hori T, Ayusawa D, Shimizu K, et al. Chromosome breakage induced by thymidylate stress in thymidylate synthase-negative mutants of mouse FM3A cells. Cancer Res 1984;44:703–709.

214. Borchers AH, Kennedy KA, Straw JA. Inhibition of DNA excision repair by methotrexate in Chinese hamster ovary cells following exposure to ultraviolet irradiation or ethylmethanesulfonate. Cancer Res 1990;50:1786–1789.

215. Goulian M, Bleile B, Tseng BY. Methotrexate-induced misincorporation of uracil into DNA. Proc Natl Acad Sci U S A 1980;77: 1956–1960.

216. Grafstrom RH, Tseng BY, Goulian M. The incorporation of uracil into animal cell DNA in vitro. Cell 1978;15:131–140.

217. Curtin NJ, Harris AL, Aherne GW. Mechanism of cell death following thymidylate synthase inhibition: 2′-deoxyuridine-5′-triphosphate accumulation, DNA damage, and growth inhibition following exposure to CB3717 and dipyridamole. Cancer Res 1991;51:2346–2352.

218. Beck WR, Wright GE, Nusbaum NJ, et al. Enhancement of methotrexate cytotoxicity by uracil analogues that inhibit deoxyuridine triphosphate nucleotidohydrolase (dUTPase) activity. Adv Exp Med Biol 1986;195:97–104.

219. Bertino JR, Goker E, Gorlick R, et al. Resistance mechanisms to methotrexate in tumors. Oncologist 1996;1:223–226.

220. Li W, Fan J, Hochhauser D, et al. Lack of functional retinoblastoma protein mediates increased resistance to antimetabolites in human sarcoma cell lines. Proc Natl Acad Sci U S A 1995;92: 10436–10440.

221. Goker E, Waltham M, Kheradpour A, et al. Amplification of the dihydrofolate reductase gene is a mechanism of acquired resistance to methotrexate in patients with acute lymphoblastic leukemia and is correlated with p53 gene mutations. Blood 1995;86:677–684.

222. Li WW, Fan J, Hochhauser D, et al. Overexpression of p21waf1 leads to increased inhibition of E2F-1 phosphorylation and sensitivity to anticancer drugs in retinoblastoma-negative human sarcoma cells. Cancer Res 1997;57:2193–2199.

223. Winter-Vann AM, Kamen BA, Bergo MO, et al. Targeting ras signaling through inhibition of carboxyl methylation: an unexpected property of methotrexate. Proc Natl Acad Sci USA 2003; 100(11):6529–6534.

224. Pinedo HM, Zaharko DS, Bull J, et al. The relative contribution of drug concentration and duration of exposure to mouse bone marrow toxicity during continuous methotrexate infusion. Cancer Res 1977;37:445–450.

225. Cherry LM, Hsu TC. Restitution of chromatid and isochromatid breaks induced in the G2 phase by actinomycin D. Environ Mutagen 1982;4:259–265.

226. Hryniuk WM, Bertino JR. Treatment of leukemia with large doses of methotrexate and folinic acid: clinical- biochemical correlates. J Clin Invest 1969;48:2140–2155.

227. Pinedo HM, Zaharko DS, Bull JM, et al. The reversal of methotrexate cytotoxicity to mouse bone marrow cells by leucovorin and nucleosides. Cancer Res 1976;36:4418–4424.

228. Howell SB, Mansfield SJ, Taetle R. Thymidine and hypoxanthine requirements of normal and malignant human cells for protection against methotrexate cytotoxicity. Cancer Res 1981;41:945–950.

229. Howell SB, Ensminger WD, Krishan A, et al. Thymidine rescue of high-dose methotrexate in humans. Cancer Res 1978;38: 325–330.

230. Rustum YM. High-pressure liquid chromatography. I. Quantitative separation of purine and pyrimidine nucleosides and bases. Anal Biochem 1978;90:289–299.

231. Willson JK, Fischer PH, Remick SC, et al. Methotrexate and dipyridamole combination chemotherapy based upon inhibition of nucleoside salvage in humans. Cancer Res 1989;49: 1866–1870.

232. Cole PD, Smith AK, Kamen BA. Osteosarcoma cells, resistant to methotrexate due to nucleoside and nucleobase salvage, are sensitive to nucleoside analogs. Cancer Chemother Pharmacol 2002;50(2):111–116.

233. Novelli A, Mini E, Liuffi M, et al. Clinical data on rescue of high-dose methotrexate with N5-methyltetrahydrofolate in human solid tumors. In: Periti P, ed. High-Dose Methotrexate Pharmacology, Toxicology and Chemotherapy. Firenze, Italy: Giuntina, 1978:299.

234. Matherly LH, Barlowe CK, Goldman ID. Antifolate polyglutamylation and competitive drug displacement at dihydrofolate reductase as important elements in leucovorin rescue in L1210 cells. Cancer Res 1986;46:588–593.

235. Bernard S, Etienne MC, Fischel JL, et al. Critical factors for the reversal of methotrexate cytotoxicity by folinic acid. Br J Cancer 1991;63:303–307.

236. Browman GP, Goodyear MD, Levine MN, et al. Modulation of the antitumor effect of methotrexate by low-dose leucovorin in squamous cell head and neck cancer: a randomized placebo- controlled clinical trial. J Clin Oncol 1990;8:203–208.

237. Yap AK, Luscombe DK. Rapid and inexpensive enzyme inhibition assay of methotrexate. J Pharmacol Methods 1986;16: 139–150.

238. Myers CE, Lippman ME, Elliot HM, et al. Competitive protein binding assay for methotrexate. Proc Natl Acad Sci U S A 1975;72:3683–3686.

239. Hande K, Gober J, Fletcher R. Trimethoprim interferes with serum methotrexate assay by the competitive protein binding technique. Clin Chem 1980;26:1617–1619.

240. Pesce MA, Bodourian SH. Evaluation of a fluorescence polarization immunoassay procedure for quantitation of methotrexate. Ther Drug Monit 1986;8:115–121.

241. Donehower RC, Hande KR, Drake JC, et al. Presence of 2,4-diamino-N10-methylpteroic acid after high-dose methotrexate. Clin Pharmacol Ther 1979;26:63–72.

242. Oellerich M, Engelhardt P, Schaadt M, et al. Determination of methotrexate in serum by a rapid, fully mechanized enzyme immunoassay (EMIT). J Clin Chem Clin Biochem 1980;18:169–174.

243. Allegra CJ, Drake JC, Bell BA, et al. Measuring levels of methotrexate [letter]. N Engl J Med 1985;313:184.

244. So N, Chandra DP, Alexander IS, et al. Determination of serum methotrexate and 7-hydroxymethotrexate concentrations. Method evaluation showing advantages of high-performance liquid chromatography. J Chromatogr 1985;337:81–90.

245. Stout M, Ravindranath Y, Kauffman R. High-performance liquid chromatographic assay for methotrexate utilizing a cold acetonitrile purification and separation of plasma or cerebrospinal fluid. J Chromatogr 1985;342:424–430.

246. Palmisano F, Cataldi TR, Zambonin PG. Determination of the antineoplastic agent methotrexate in body fluids by high- performance liquid chromatography with electrochemical detection. J Chromatogr 1985;344:249–258.

247. Slordal L, Prytz PS, Pettersen I, et al. Methotrexate measurements in plasma: comparison of enzyme multiplied immunoassay technique, TDx fluorescence polarization immunoassay, and high pressure liquid chromatography. Ther Drug Monit 1986;8:368–372.

248. Zaharko DS, Dedrick RL, Bischoff KB, et al. Methotrexate tissue distribution: prediction by a mathematical model. J Natl Cancer Inst 1971;46:775–784.

249. Chungi VS, Bourne DW, Dittert LW. Drug absorption VIII: kinetics of GI absorption of methotrexate. J Pharm Sci 1978;67: 560–561.

250. Stuart JF, Calman KC, Watters J, et al. Bioavailability of methotrexate: implications for clinical use. Cancer Chemother Pharmacol 1979;3:239–241.

251. Balis FM, Savitch JL, Bleyer WA. Pharmacokinetics of oral methotrexate in children. Cancer Res 1983;43:2342–2345.

252. Balis FM, Holcenberg JS, Poplack DG, et al. Pharmacokinetics and pharmacodynamics of oral methotrexate and mercaptopurine in children with lower risk acute lymphoblastic leukemia: a joint children's cancer group and pediatric oncology branch study. Blood 1998;92:3569–3577.

253. Steele WH, Lawrence JR, Stuart JF, et al. The protein binding of methotrexate by the serum of normal subjects. Eur J Clin Pharmacol 1979;15:363–366.

254. Liegler DG, Henderson ES, Hahn MA, et al. The effect of organic acids on renal clearance of methotrexate in man. Clin Pharmacol Ther 1969;10:849–857.

255. Wan SH, Huffman DH, Azarnoff DL, et al. Effect of route of administration and effusions on methotrexate pharmacokinetics. Cancer Res 1974;4:3487–3491.

256. Chabner BA, Stoller RG, Hande K, et al. Methotrexate disposition in humans: case studies in ovarian cancer and following high-dose infusion. Drug Metab Rev 1978;8:107–117.

257. Torres IJ, Litterst CL, Guarino AM. Transport of model compounds across the peritoneal membrane in the rat. Pharmacology 1978;17:330–340.

258. Fossa SD, Heilo A, Bormer O. Unexpectedly high serum methotrexate levels in cystectomized bladder cancer patients with an ileal conduit treated with intermediate doses of the drug. J Urol 1990;143:498–501.

259. Kristenson L, Weismann K, Hutters L. Renal function and the rate of disappearance of methotrexate from serum. Eur J Clin Pharmacol 1975;8:439–444.

260. Bressolle F, Bologna C, Kinowski JM, et al. Effects of moderate renal insufficiency on pharmacokinetics of methotrexate in rheumatoid arthritis patients. Ann Rheum Dis 1998;57:110–113.

261. Bleyer WA. The clinical pharmacology of methotrexate: new applications of an old drug. Cancer 1978;41:36–51.

262. Howell SB, Tamerius RK. Achievement of long duration methotrexate exposure with concurrent low dose thymidine protection: influence of methotrexate pharmacokinetics. Eur J Cancer 1980;16:1427–1432.

263. Evans WE, Crom WR, Stewart CF, et al. Methotrexate systemic clearance influences probability of relapse in children with standard-risk acute lymphocytic leukaemia. Lancet 1984;1:359–362.

264. Evans WE, Crom WR, Abromowitch M, et al. Clinical pharmacodynamics of high-dose methotrexate in acute lymphocytic leukemia. Identification of a relation between concentration and effect. N Engl J Med 1986;314:471–477.

265. Borsi JD, Moe PJ. Systemic clearance of methotrexate in the prognosis of acute lymphoblastic leukemia in children. Cancer 1987;60:3020–3024.

266. Pearson AD, Amineddine HA, Yule M, et al. The influence of serum methotrexate concentrations and drug dosage on outcome in childhood acute lymphoblastic leukaemia [see comments]. Br J Cancer 1991;64:169–173.

267. Calvert AH, Bondy PK, Harrap KR. Some observations on the human pharmacology of methotrexate. Cancer Treat Rep 1977;61:1647–1656.

268. Monjanel S, Rigault JP, Cano JP, et al. High-dose methotrexate: preliminary evaluation of a pharmacokinetic approach. Cancer Chemother Pharmacol 1979;3:189–196.

269. Sasaki K, Tanaka J, Fujimoto T. Theoretically required urinary flow during high-dose methotrexate infusion. Cancer Chemother Pharmacol 1984;13:9–13.

270. Romolo JL, Goldberg NH, Hande KR, et al. Effect of hydration on plasma-methotrexate levels. Cancer Treat Rep 1977;61: 1393–1396.

271. Huang KC, Wenczak BA, Liu YK. Renal tubular transport of methotrexate in the rhesus monkey and dog. Cancer Res 1979;39:4843–4848.

272. Iven H, Brasch H. Influence of the antibiotics piperacillin, doxycycline, and tobramycin on the pharmacokinetics of methotrexate in rabbits. Cancer Chemother Pharmacol 1986;17:218–222.

273. Iven H, Brasch H. The effects of antibiotics and uricosuric drugs on the renal elimination of methotrexate and 7- hydroxymethotrexate in rabbits. Cancer Chemother Pharmacol 1988;21:337–342.

274. Iven H, Brasch H. Cephalosporins increase the renal clearance of methotrexate and 7-hydroxymethotrexate in rabbits. Cancer Chemother Pharmacol 1990;26:139–143.

275. Titier K, Lagrange F, Pehourcq F, et al. Pharmacokinetic interaction between high-dose methotrexate and oxacillin. Ther Drug Monit 2002;24(4):570–572.

276. Dalle JH, Auvrignon A, Vassal G etal. Interaction between methotrexate and ciprofloxacin. J Pediatr Hematol Oncol 2002; 4(4):321–322.

277. Kerr IG, Jolivet J, Collins JM, et al. Test dose for predicting high-dose methotrexate infusions. Clin Pharmacol Ther 1983;33: 44–51.

278. Favre R, Monjanel S, Alfonsi M, et al. High-dose methotrexate: a clinical and pharmacokinetic evaluation. Treatment of advanced squamous cell carcinoma of the head and neck using a prospective mathematical model and pharmacokinetic surveillance. Cancer Chemother Pharmacol 1982;9:156–160.

279. Gewirt DA, White JC, Goldman ID. Transport, binding and polyglutamation of methotrexate (MTX) in freshly isolated hepatocytes. Am Assoc Cancer Res 1979:147.

280. Strum WB, Liem HH. Hepatic uptake, intracellular protein binding and biliary excretion of amethopterin. Biochem Pharmacol 1977;26:1235–1240.

281. Jacobs SA, Derr CJ, Johns DG. Accumulation of methotrexate diglutamate in human liver during methotrexate therapy. Biochem Pharmacol 1977;26:2310–2313.

282. Strum WB, Liem HH, Muller-Eberhard U. Effect of chemotherapeutic agents on the uptake and excretion of amethopterin by the isolated perfused rat liver. Cancer Res 1978;38:4734–4736.

283. Said HM, Hollander D. Inhibitory effect of bile salts on the enterohepatic circulation of methotrexate in the unanesthetized rat: inhibition of methotrexate intestinal absorption. Cancer Chemother Pharmacol 1986;16:121–124.

284. Lerne PR, Creaven PJ, Allen LM, et al. Kinetic model for the disposition and metabolism of moderate and high-dose methotrexate in man. Cancer Chemother Rep 1975;59:811–817.

285. Shen DD, Azarnoff DL. Clinical pharmacokinetics of methotrexate. Clin Pharmacokinet 1978;3:1–13.

286. Steinberg SE, Campbell CL, Bleyer WA, et al. Enterohepatic circulation of methotrexate in rats in vivo. Cancer Res 1982;42:1279–1282.

287. Breithaupt H, Kuenzlen E. Pharmacokinetics of methotrexate and 7-hydroxymethotrexate following infusions of high- dose methotrexate. Cancer Treat Rep 1982;66:1733–1741.

288. Erttmann R, Landbeck G. Effect of oral cholestyramine on the elimination of high-dose methotrexate. J Cancer Res Clin Oncol 1985;110:48–50.

289. Jacobs SA, Stoller RG, Chabner BA, et al. 7-Hydroxymethotrexate as a urinary metabolite in human subjects and rhesus monkeys receiving high dose methotrexate. J Clin Invest 1976;57:534–538.

290. Bremnes RM, Slordal L, Wist E, et al. Formation and elimination of 7-hydroxymethotrexate in the rat in vivo after methotrexate administration. Cancer Res 1989;49:2460–2464.

291. Sholar PW, Baram J, Seither R, et al. Inhibition of folate- dependent enzymes by 7-OH-methotrexate. Biochem Pharmacol 1988;37:3531–3534.

292. Clendeninn NJ, Drake JC, Allegra CJ, et al. Methotrexate polyglutamates have a greater affinity and more rapid on- rate for purified human dihydrofolate reductase than MTX. Proc Am Assoc Cancer Res 1985:232.

293. McCullough JL, Chabner BA, Bertino JR. Purification and properties of carboxypeptidase G1. J Biol Chem 1971;246: 7207–7213.

294. Abelson HT, Ensminger W, Rosowsky A, et al. Comparative effects of citrovorum factor and carboxypeptidase G1 on cerebrospinal fluid-methotrexate pharmacokinetics. Cancer Treat Rep 1978;62:1549–1552.

295. Chabner BA, Young RC. Threshold methotrexate concentration for in vivo inhibition of DNA synthesis in normal and tumorous target tissues. J Clin Invest 1973;52:1804–1811.

296. Sirotnak FM, Moccio DM. Pharmacokinetic basis for differences in methotrexate sensitivity of normal proliferative tissues in the mouse. Cancer Res 1980;40:1230–1234.

297. Toffoli G, Russo A, Innocenti F etal. Effect of methylenetetrahydrofolate reductase C677T polymorphism on toxicity and homocysteine plasma level after chronic methotrexate treatment of ovarian cancer patients. Int J Cancer 2003;103(3):294-299.

298. Chiusolo P, Reddiconto G, Casorelli I, et al. Preponderance of methylenethtrahydrofolate reductase C677T homozygosity among leukemia patients intolerant to methotrexate. Ann Oncol 2002;13(12):1915–1918.

299. Ulrich CM, Yasui Y, Storb R, et al. Pharmacogenetics of methotrexate: toxicity among marrow transplantation patients varies with the methylenetetrahydrofolate reductase C677T polymorphism. Blood 2001;98(1):231–234.

300. Urano W, Taniguchi A, Yamanaka H, et al. Polymorphisms in the methylenetetrahydrofolate reductase gene were associated with both the efficacy and the toxicity of methotrexate used for the treatment of rheumatoid arthritis, as evidenced by single locus haplotype analysis. Pharmacogenetics 2002;12(3):183–190.

301. Van Ede AE, Laan RF, Blom HJ, et al. The C677T mutation in the methylenetetrahydrofolate reductase gene: a genetic risk factor for methotrexate-related elevation of liver enzymes in rheumatoid arthritis patients. Arthritis Rheum 2001;44(11):2525–2530.

302. Ackland SP, Schilsky RL. High-dose methotrexate: a critical reappraisal. J Clin Oncol 1987;5:2017–2031.

303. Von Hoff DD, Penta JS, Helman LJ, et al. Incidence of drug- related deaths secondary to high-dose methotrexate and citrovorum factor administration. Cancer Treat Rep 1977;61:745–748.

304. Hempel L, Misselwitz J, Fleck C etal. Influence of high-dose methotrexate therapy on glomerular and tubular kidney function. Med Pediatr Oncol 2003;40(6):348–354.

305. Stoller RG, Hande KR, Jacobs SA, et al. Use of plasma pharmacokinetics to predict and prevent methotrexate toxicity. N Engl J Med 1977;297:630–634.

306. Bacci G, Ferrari S, Longhi A, et al. Delayed methotrexate clearance in osteosarcoma patients treated with multiagent regimens of neoadjuvant chemotherapy. Oncol Rep 2003;10(4):851–857.

307. Thyss A, Milano G, Kubar J, et al. Clinical and pharmacokinetic evidence of a life-threatening interaction between methotrexate and ketoprofen. Lancet 1986;1:256–258.

308. Takeda M, Khamdang S, Narikawa S, et al. Characterization of methotrexate transport and its drug interactions with human organic anion transporters. J Pharmacol Exp Ther 2002;302(2): 666–671.

309. Nozaki Y, Kusuhara H, Endou H, et al. Quantitative evaluation of the drug-drug interactions between methotrexate and nonsteroidal anti-inflammatory drugs in the renal uptake process based on the contribution of organic anion transporters and reduced folate carrier. J Pharmacol Exp Ther 2004;309(1):226–234.

309a. Wall, S.M., Johansen, M.J., Molony, D.A., DuBose, T.D., Jr., Jaffe, N., and Madden, T. Effective clearance of methotrexate using high-flux hemodialysis membranes. Am J Kidney Dis 1996;28:846–854.

310. Bertino JR, Condos S, Horvath C, et al. Immobilized carboxypeptidase G1 in methotrexate removal. Cancer Res 1978;38:1936–1941.

311. DeAngelis LM, Tong WP, Lin S, et al. Carboxypeptidase G2 rescue after high-dose methotrexate. J Clin Oncol 1996;14: 2145–2149.

312. Mohty M, Peyriere H, Guinet C, et al. Carboxypeptidase G2 rescue in delayed methotrexate elimination in renal failure. Leuk Lymphoma 2000;37(3-4):441–443.

313. Krause AS, Weihrauch MR, Bode U, et al. Carboxypeptidase-G2 rescue in cancer patients with delayed methotrexate elimination after high-dose methotrexate therapy. Leuk Lymphoma 2002;43(11):2139–2143.

314. Widemann BC, Balis FM, Murphy RF, et al. Carboxypeptidase-G2, thymidine, and leucovorin rescue in cancer patients with methotrexate-induced renal dysfunction. J Clin Oncol 1997;15: 2125–2134.

315. Schornagel JH, Leyva A, Bucsa JM, et al. Thymidine prevention of methotrexate toxicity in head-and-neck cancer. In Pinedo HM, ed. Clinical Pharmacology of Antineoplastic Drugs. Amsterdam: Elsevier/North Holland, 1978:83.

316. Van den Bongard HJ, Mathjt RA, Boogerd W, et al. Successful rescue with leucovorin and thymidine in a patient with high-dose methotrexate induced acute renal failure. Cancer Chemother Pharmacol 2001;47(6):537–540.

317. Howell SB, Herbst K, Boss GR, et al. Thymidine requirements for the rescue of patients treated with high-dose methotrexate. Cancer Res 1980;40:1824–1829.

318. Capizzi RL. Schedule-dependent synergism and antagonism between methotrexate and L-asparaginase. Biochem Pharmacol 1974;23:151.

319. Yap BS, McCredie KB, Benjamin RS, et al. Refractory acute leukaemia in adults treated with sequential colaspase and high-dose methotrexate. BMJ 1978;2:791–793.

320. Moran RG, Mulkins M, Heidelberger C. Role of thymidylate synthetase activity in development of methotrexate cytotoxicity. Proc Natl Acad Sci U S A 1979;76:5924–5928.

321. Rajeswari R, Shetty PA, Gothoskar BP, et al. Pharmacokinetics of methotrexate in adult Indian patients and its relationship to nutritional status. Cancer Treat Rep 1984;68:727–732.

322. Mihranian MH, Wang YM, Daly JM. Effects of nutritional depletion and repletion on plasma methotrexate pharmacokinetics. Cancer 1984;54:2268–2271.

323. Zachariae H, Kragballe K, Sogaard H. Methotrexate induced liver cirrhosis. Studies including serial liver biopsies during continued treatment. Br J Dermatol 1980;102:407–412.

324. Dahl MG, Gregory MM, Scheuer PJ. Methotrexate hepatotoxicity in psoriasis—comparison of different dose regimens. BMJ 1972;1:654–656.

325. Podurgiel BJ, McGill DB, Ludwig J, et al. Liver injury associated with methotrexate therapy for psoriasis. Mayo Clin Proc 1973;48:787–792.

326. Willkens RF, Clegg DO, Ward JR, et al. Liver biopsies in patients on low-dose pulse methotrexate for the treatment of rheumatoid arthritis [abstract]. In: Sixteenth International Congress on Rheumatology. Sydney, Australia: 1985:88.

327. Mackenzie AH. Hepatotoxicity of prolonged methotrexate therapy for rheumatoid arthritis. Cleve Clin Q 1985;52:129–135.

328. Scully CJ, Anderson CJ, Cannon GW. Long-term methotrexate therapy for rheumatoid arthritis. Semin Arthritis Rheum 1991;20:317–331.

329. Phillips CA, Cera PJ, Mangan TF, et al. Clinical liver disease in patients with rheumatoid arthritis taking methotrexate. J Rheumatol 1992;19:229–233.

330. Weber BL, Tanyer G, Poplack DG, et al. Transient acute hepatotoxicity of high-dose methotrexate therapy during childhood. J Natl Cancer Inst Monogr 1987;5:207–212.

331. Clarysse AM, Cathey WJ, Cartwright GE, et al. Pulmonary disease complicating intermittent therapy with methotrexate. JAMA 1969;209:1861–1868.

332. Sostman HD, Matthay RA, Putman CE, et al. Methotrexate- induced pneumonitis. Medicine (Baltimore)1976;55:371–388.

333. Akoun GM, Mayaud CM, Touboul JL, et al. Use of bronchoalveolar lavage in the evaluation of methotrexate lung disease. Thorax 1987;42:652–655.

334. Kremer JM, Phelps CT. Long-term prospective study of the use of methotrexate in the treatment of rheumatoid arthritis. Update after a mean of 90 months. Arthritis Rheum 1992;35:138–145.

335. Searles G, McKendry RJ. Methotrexate pneumonitis in rheumatoid arthritis: potential risk factors. Four case reports and a review of the literature. J Rheumatol 1987;14:1164–1171.

336. Carson CW, Cannon GW, Egger MJ, et al. Pulmonary disease during the treatment of rheumatoid arthritis with low dose pulse methotrexate. Semin Arthritis Rheum 1987;16:186–195.

337. Kremer JM, Alarcon GS, Weinblatt ME, et al. Clinical, laboratory, radiographic, and histopathologic features of methotrexate-associated lung injury in patients with rheumatoid arthritis: a multicenter study with literature review [see comments]. Arthritis Rheum 1997;40:1829–1837.

338. Hargreaves MR, Mowat AG, Benson MK. Acute pneumonitis associated with low dose methotrexate treatment for rheumatoid arthritis: report of five cases and review of published reports. Thorax 1992;47:628–633.

339. Alarcon GS, Kremer JM, Macaluso M, et al. Risk factors for methotrexate-induced lung injury in patients with rheumatoid arthritis. A multicenter, case-control study. Methotrexate Lung Study Group. Ann Intern Med 1997;127:356–364.

340. Goldberg NH, Romolo JL, Austin EH, et al. Anaphylactoid type reactions in two patients receiving high dose intravenous methotrexate. Cancer 1978;41:52–55.

341. Doyle LA, Berg C, Bottino G, et al. Erythema and desquamation after high-dose methotrexate. Ann Intern Med 1983;98: 611–612.

342. Shamberger RC, Rosenberg SA, Seipp CA, et al. Effects of high-dose methotrexate and vincristine on ovarian and testicular functions in patients undergoing postoperative adjuvant treatment of osteosarcoma. Cancer Treat Rep 1981;65:739–746.

343. Shapiro WR, Young DF, Mehta BM. Methotrexate: distribution in cerebrospinal fluid after intravenous, ventricular and lumbar injections. N Engl J Med 1975;293:161–166.

344. Tatef ML, MargolinKA, Doroshow JH, et al. Pharmacokinetics and toxicity of high-dose intravenous methotrexate in the treatment of leptomeningeal carcinomatosis. Cancer Chemother Pharmacol 2000;46(1):19–26.

345. Bleyer WA, Drake JC, Chabner BA. Neurotoxicity and elevated cerebrospinal-fluid methotrexate concentration in meningeal leukemia. N Engl J Med 1973;289:770–773.

346. Morse M, Savitch J, Balis F, et al. Altered central nervous system pharmacology of methotrexate in childhood leukemia: another sign of meningeal relapse. J Clin Oncol 1985;3:19–24.

347. Blaney SM, Balis FM, Poplack DG. Current pharmacological treatment approaches to central nervous system leukaemia. Drugs 1991;41:702–716.

348. Ettinger LJ, Chervinsky DS, Freeman AI, et al. Pharmacokinetics of methotrexate following intravenous and intraventricular administration in acute lymphocytic leukemia and non-Hodgkin's lymphoma. Cancer 1982;50:1676–1682.

349. Grossman SA, Reinhard CS, Loats HL. The intracerebral penetration of intraventricularly administered methotrexate: a quantitative autoradiographic study. J Neurooncol 1989;7:319–328.

350. Bleyer WA, Poplack DG, Simon RM. “Concentration ? time” methotrexate via a subcutaneous reservoir: a less toxic regimen for intraventricular chemotherapy of central nervous system neoplasms. Blood 1978;51:835–842.

351. Glantz MJ, Cole BF, Recht L, et al. High-dose intravenous methotrexate for patients with nonleukemic leptomeningeal cancer: is intrathecal chemotherapy necessary? J Clin Oncol 1998;16:1561–1567.

352. Peylan-Ramu N, Poplack DG, Blei CL, et al. Computer assisted tomography in methotrexate encephalopathy. J Comput Assist Tomogr 1977;1:216–221.

353. Paakko E, Vainionpaa L, Lanning M, et al. White matter changes in children treated for acute lymphoblastic leukemia. Cancer 1992;70:2728–2733.

354. Shapiro WR, Allen JC, Horten BC. Chronic methotrexate toxicity to the central nervous system. Clin Bull 1980;10:49–52.

355. Jaffe N, Takaue Y, Anzai T, et al. Transient neurologic disturbances induced by high-dose methotrexate treatment. Cancer 1985;56:1356–1360.

356. Fritsch G, Urban C. Transient encephalopathy during the late course of treatment with high-dose methotrexate. Cancer 1984; 53:1849–1851.

357. Walker RW, Allen JC, Rosen G, et al. Transient cerebral dysfunction secondary to high-dose methotrexate. J Clin Oncol 1986; 4:1845–1850.

358. Kubo M, Azuma E, Arai S, et al. Transient encephalopathy following a single exposure of high-dose methotrexate in a child with acute lymphoblastic leukemia. Pediatr Hematol Oncol 1992;9:157–165.

359. Allen J, Rosen G, Juergens H, et al. The inability of oral leucovorin to elevate CSF 5-methyl-tetrahydrofolate following high dose intravenous methotrexate therapy. J Neurooncol 1983; 1:39–44.

360. Mehta BM, Glass JP, Shapiro WR. Serum and cerebrospinal fluid distribution of 5-methyltetrahydrofolate after intravenous calcium leucovorin and intra-Ommaya methotrexate administration in patients with meningeal carcinomatosis. Cancer Res 1983;43:435–438.

361. Ochs J, Mulhern R, Fairclough D, et al. Comparison of neuropsychologic functioning and clinical indicators of neurotoxicity in long-term survivors of childhood leukemia given cranial radiation or parenteral methotrexate: a prospective study. J Clin Oncol 1991;9:145–151.

362. Mahoney DH Jr, Shuster JJ, Nitschke R, et al. Acute neurotoxicity in children with B-precursor acute lymphoid leukemia: an association with intermediate-dose intravenous methotrexate and intrathecal triple therapy—a Pediatric Oncology Group study. J Clin Oncol 1998;16:1712–1722.

363. Phillips PC, Dhawan V, Strother SC, et al. Reduced cerebral glucose metabolism and increased brain capillary permeability following high-dose methotrexate chemotherapy: a positron emission tomographic study. Ann Neurol 1987;21:59–63.

364. Kishi S, Griener J, Cheng C, et al. Homocysteine, pharmacogenetics, and neurotoxicity in children with leukemia. J Clin Oncol 2003;21(16):3084–3091.

365. Gilbert MR, Harding BL, Grossman SA. Methotrexate neurotoxicity: in vitro studies using cerebellar explants from rats. Cancer Res 1989;49:2502–2505.

366. Livrea P, Trojano M, Simone IL, et al. Acute changes in blood- CSF barrier permselectivity to serum proteins after intrathecal methotrexate and CNS irradiation. J Neurol 1985;231:336–339.

367. Spiegel RJ, Cooper PR, Blum RH, et al. Treatment of massive intrathecal methotrexate overdose by ventriculolumbar perfusion. N Engl J Med 1984;311:386–388.

368. Shih C, Chen VJ, Gossett LS, et al. LY231514, a pyrrolo[2,3- d]pyrimidine-based antifolate that inhibits multiple folate- requiring enzymes. Cancer Res 1997;57:1116–1123.

368a. Wang Y, Zhao R, Chattopadhyay S, et al. A novel folate transport activity in human mesothelioma cell lines with high affinity and specificity for the new-generation antifolate, Pemetrexed. Cancer Res 2002, 62:6434–6437.

369. Shih C, Habeck LL, Mendelsohn LG, et al. Multiple folate enzyme inhibition: mechanism of a novel pyrrolopyrimidine-based antifolate LY231514 (MTA). Adv Enzyme Regul 1998; 38:135–152.

370. Worzalla JF, Shih C, Schultz RM. Role of folic acid in modulating the toxicity and efficacy of the multitargeted antifolate, LY231514. Anticancer Res 1998;18:3235–3239.

371. Scagliotti GV, Shin DM, Kindler HL, et al. Phase II study of pemetrexed with and without folic acid and vitamin B12 as front-line therapy in malignant pleural mesothelioma. J Clin Oncol 2003;21(8):1556–1561.

372. Vogelzang NJ, Rusthoven JJ, Symanowski J, et al. Phase III study of pemetrexed in combination with cisplatin versus cisplatin alone in patients with malignant pleural mesothelioma. J Clin Oncol 2003;21(14):2636–2644

373. Clarke SJ, Abatt R, Goedhals L, et al. Phase II trial of pemetrexed disodium (Alimta, LY231514) in chemotherapy-naive patients with advanced non-small-cell lung cancer. Ann Oncol 2002;13(5):737–741.

374. Hanna N, Shepherd FA, Fossella FV, et al. Randomized phase III trial of pemetrexed versus docetaxel in patients with non-small-cell lung cancer previously treated with chemotherapy. J Clin Oncol 2004;22(9):1589–1597.

375. Rafi I, Taylor GA, Calvete JA, et al. Clinical pharmacokinetic and pharmacodynamic studies with the nonclassical antifolate thymidylate synthase inhibitor 3, 4-dihydro-2- amino-6-methyl-4-oxo-5-(4-pyridylthio)-quinazoline dihydrochloride (AG337) given by 24-hour continuous intravenous infusion. Clin Cancer Res 1995;1:1275–1284.

376. Webber S, Bartlett CA, Boritzki TJ, et al. AG337, a novel lipophilic thymidylate synthase inhibitor: in vitro and in vivo preclinical studies. Cancer Chemother Pharmacol 1996;37: 509–517.

377. Creaven PJ, Pendyala L, Meropol NJ, et al. Initial clinical trial and pharmacokinetics of Thymitaq (AG337) by 10-day continuous infusion in patients with advanced solid tumors. Cancer Chemother Pharmacol 1998;41:167–170.

378. Hughes AN, Rafi I, Griffin MJ, et al. Phase I studies with the nonclassical antifolate nolatrexed dihydrochloride (AG337, THYMITAQ) administered orally for 5 days. Clin Cancer Res 1999;5:111–118.

379. Pivot X, Wadler S, Kelly C, et al. Result of two randomized trials comparing nolatrexed (Thymitaq) versus methotrexate in patients with recurrent head and neck cancer. Ann Oncol 2001; 12(11):1595–1599.

380. Zalcberg JR, Cunningham D, Van Cutsem E, et al. ZD1694: a novel thymidylate synthase inhibitor with substantial activity in the treatment of patients with advanced colorectal cancer. Tomudex Colorectal Study Group. J Clin Oncol 1996;14: 716–721.

381. Cunningham D, Zalcberg J, Smith I, et al. “Tomudex” (ZD1694): a novel thymidylate synthase inhibitor with clinical antitumour activity in a range of solid tumours. “Tomudex” International Study Group. Ann Oncol 1996;7:179–182.

382. Cunningham D. Mature results from three large controlled studies with raltitrexed (“Tomudex”). Br J Cancer 1998;77:15–21.

383. Van Cutsem E, Cunningham D, Maroun J, et al. Raltitrexed: current clinical status and future directions. Ann Oncol 2002; 13(4):513–522.

384. Yin MB, Guo B, Panadero A, et al. Cyclin E-cdk2 activation is associated with cell cycle arrest and inhibition of DNA replication induced by the thymidylate synthase inhibitor Tomudex. Exp Cell Res 1999;247:189–199.

385. Santini D, Massacesi C, D'Angelillo RM, et al. Raltitrexed plus weekly oxaliplatin as a first-line chemotherapy in metastatic colorectal cancer: a multicenter non-randomized phase II study. Med Oncol 2004;21(1):59–66.

386. Feliu J, Salud A, Escudero P, et al. Irinotecan plus raltitrexed as first-line treatment in advanced colorectal cancer: a phase II study. Br J Cancer 2003;90(8):1502–1507.

387. Farrugia DC, Ford HE, Cunningham D, et al. Thymidylate synthase expression in advanced colorectal cancer predicts for response to raltitrexed. Clin Cancer Res 2003;9(2):792–801.

388. Hainsworth J, Vergote I, Janssens J. A review of phase II studies of ZD9331 treatment for relapsed or refractory solid tumours. Anticancer Drugs 2003;14(Suppl1):S13–19.

389. Springer CJ, Poon GK, Sharma SK, et al. Analysis of antibody-enzyme conjugate clearance by investigation of prodrug and active drug in an ADEPT clinical study. Cell Biophys 1994;25: 193–207.

390. Springer CJ, Bavetsias V, Jackman AL, et al. Prodrugs of thymidylate synthase inhibitors: potential for antibody directed enzyme prodrug therapy (ADEPT). Anticancer Drug Des 1996;11:625–636.

391. Syrigos KN, Epenetos AA. Antibody directed enzyme prodrug therapy (ADEPT): a review of the experimental and clinical considerations. Anticancer Res 1999;19:605–613.