Farnesol Synthesis Essay

Department of Biological Sciences, Florida International University, Miami, FL 33199, USA

Copyright © 2014 Fernando G. Noriega. This is an open access article distributed under the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

Our understanding of JH biosynthesis has significantly changed in the last years. In this review I would like to discuss the following topics: (1) the progresses in understanding the JH biosynthesis pathway. Access to genome sequences has facilitated the identification of all the genes encoding biosynthetic enzymes and the completion of comprehensive transcriptional studies, as well as the expression and characterization of recombinant enzymes. Now the existence of different flux directionalites, feed-back loops and pathway branching points in the JH biosynthesis pathways can be explored; (2) the new concepts in the modulation of JH synthesis by allatoregulators. The list of putative JH modulators is increasing. I will discuss their possible role during the different physiological states of the CA; (3) the new theoretical and physiological frameworks for JH synthesis analysis. I will discuss the bases of the flux model for JH biosynthesis. JH plays multiple roles in the control of ovary development in female mosquitoes; therefore, the CA presents different physiological states, where JH synthesis is altered by gating the flux at distinctive points in the pathway; (4) in the final section I will identify new challenges and future directions on JH synthesis research.

1. Introduction

Juvenile hormone (JH) regulates development and reproductive maturation in insects [1, 2]; therefore, interruption of JH biosynthesis has been considered as a strategy for the development of target-specific insecticides [3]. Although degradation plays a role, JH titer is primarily determined by the rate of biosynthesis in the corpora allata gland (CA). A number of recent reviews have summarized the current knowledge on JH biosynthesis in insects [1, 2], as well as its potential as a target for insecticide discovery [3]. In the present review I would like to focus on the discussion of some new advances in the field and on the identification of outstanding questions that remain to be addressed, as well as the potential directions for future research.

Our understanding of JH biosynthesis has significantly changed over the past few years. Access to genome sequences has facilitated the identification of all the genes encoding JH biosynthetic enzymes [4–6] and the completion of comprehensive transcriptional studies [5, 6], as well as the expression and characterization of recombinant JH biosynthetic enzymes [7–10]. The development of new technologies is facilitating the analysis of JH biosynthesis rates, enzymatic activities, and metabolite pool sizes in the CA [11, 12]. In addition, new theoretical and physiological frameworks are simplifying JH synthesis analysis [12].

This review will emphasize the work that has been done on the biosynthesis of JH III in the mosquito Aedes aegypti. The importance of A. aegypti as a vector of diseases has attracted the interest of scientists and funding agencies for many years. Consequently, there is ample information available on biological, ecological, anatomical, and physiological aspects of this mosquito, both published in primary research articles and summarized in excellent textbooks [13, 14]. Vectorbase is an excellent web resource available for genomic analysis [15, 16]. Molecular tools such as DNA microarrays [17, 18], RNA interference (RNAi) [10, 19], generation of transgenic lines [20, 21], and high throughput transcript sequencing approaches [22] are readily available. All these factors have contributed to make A. aegypti an excellent model for the study of JH biosynthesis.

2. JH Structures, Functions, and Mode of Action

JHs are lipophilic molecules commonly produced and released into the hemolymph by the CA, generally a pair of endocrine glands connected to the brain [24]. The naturally occurring JHs are a family of acyclic sesquiterpenoids primarily limited to insects. Eight different forms of JH have been identified. JH III is found in the majority of insects studied [2, 25], including A. aegypti [11, 25, 26]. Five JHs have been reported in Lepidoptera: JH 0, JH I, JH II, JH III, and 4-methyl JH I [27–30]. In addition, Drosophila melanogaster CA secretes a bis-epoxide JH III (JHB III) [31], as well as methyl farnesoate (MF) [32–36]. Recently, another bis-epoxide form of JH III, skipped bisepoxide (JHSB III), has been reported in heteropteran insects [37, 38]. At least one JH homologue has been identified in over 100 insect species covering more than 10 insect orders [2]. With more than 2.5 million insect species estimated to inhabit earth [39], it is reasonable to think that additional forms of JH might be discovered in the future.

The JHs are involved in reproduction, caste determination, behavior, stress response, diapause, and several polyphenisms [40]. Understanding the mode of action of JH at the molecular level has been a major challenge in insect biology. The recent discovery that the JH-resistance gene, Methoprene-tolerant (Met), plays a critical role in insect metamorphosis [41–43] has been followed by a rapid increase in our understanding of JH signaling. Met is a bHLH-PAS protein, characterized by a short stretch of basic amino acids followed by a HLH domain and two variably spaced PAS domains (A and B) [44, 45]. The idea that JH could be an activating ligand for Met was surprising because there were no examples of bHLH-PAS proteins working as hormone receptors that act as ligand-dependent transcription factors [43].

To form active transcription factors, functionally specialized bHLH-PAS proteins, such as Met, pair with a partner of their family. JH-dependent interaction between Met and its partner Taiman/SRC requires the hormone to be bound to a specific ligand-binding site. Met binds JH and its mimics with high affinity through a well-conserved hydrophobic pocket within its PAS-B domain [45]. In the absence of JH, Met is present as an inactive homodimer. Upon JH binding to the PAS-B domain, Met undergoes a conformational change that liberates Met from the homodimer complex and allows it to bind Taiman [43, 45–47]. By sensing JH and forming a ligand-dependent complex with a partner of its own bHLH-PAS family, Met establishes a unique class of intracellular hormone receptors. The complex recognizes JH-responsive elements (JHRE) in the promoter of genes containing canonical E box motifs [45, 48, 49].

In mosquitoes, JH acts via Met to regulate posteclosion development of the fat body and plays a dual role. Thousands of genes are active when the JH titer is low and then are suppressed by the rising JH; other genes appear specifically when the JH titer is high [50, 51]. Jindra et al. [43] identified some of the outstanding questions that remain unanswered after the characterization of the JH receptor. Among them I would like to highlight the following two: (1) what is the relationship among different JHs, different bHLH-PAS proteins, and diverse biological functions of JH in different systems? (2) Are different JH homologues acting in distinct ways through different complexes involving Met, Taiman, Cycle, or members of the nuclear receptor superfamily such as ultraspiracle? Recent studies in the heteropteran linden bug, Pyrrhocoris apterus, indicate that JH stimulates oogenesis through Met and Taiman but regulates gene expression in the gut through interactions of Met with the circadian proteins Clock and Cycle [52]; the latter bHLH-PAS protein has indeed been shown to bind Met in a JH-dependent manner [53]. More answers to these questions are sure to be provided in the next few years.

3. JH Biosynthetic Pathway

JH is synthesized through the mevalonate pathway (MVAP), an ancient metabolic pathway present in the three domains of life [54]. The MVAP is responsible for the synthesis of many essential molecules required for cell signaling, membrane integrity, energy homeostasis, protein prenylation, and glycosylation [55–58]. The MVAP consists of a main trunk followed by subbranches that generate a diverse range of biomolecules. Insects lack the cholesterol-synthetic branch present in vertebrates, but in the CA the MVAP branches into the synthesis of JH [59]. The biosynthetic pathway of JH III in the CA of insects involves 13 discrete enzymatic reactions and is conventionally divided into early (MVAP) and late (JH-branch) steps [2] (Figure 1).

Figure 1: JH biosynthesis pathway. The biosynthesis of JH III involves 13 enzymatic reactions that can be conventionally divided into early (MVAP) and late (JH-branch) steps. Metabolites are shown in bold and enzymes in italic. Chemical structures are in [4].

3.1. Early Steps (MVAP)

The early steps follow the MVAP to form farnesyl pyrophosphate (FPP) [59]. Initially, three units of acetyl-CoA are condensed into mevalonate by means of three sequential steps involving the enzymes acetoacetyl-CoA thiolase (THIOL), HMG-CoA synthase (HMGS), and HMG-CoA reductase (HMGR). Mevalonate is then converted to isopentenyl diphosphate (IPP) through three enzymatic reactions catalyzed by mevalonate kinase (MevK), phosphomevalonate kinase (P-MevK), and mevalonate diphosphate decarboxylase (PP-MevD) [6, 59]. FPP synthase (FPPS), a short-chain prenyltransferase, generates FPP by completing two sequential couplings: first IPP and dimethylallyl pyrophosphate (DMAPP) can condense in a head-to-tail manner to produce geranyl diphosphate (GPP). This type of head-to-tail condensation can be repeated by the further reaction of GPP with IPP yielding FPP.

FPP synthases have been identified from several insects and are typically active as homodimers [60–64]. In the mustard leaf beetle Phaedon cochleariae, FPPS possesses an interesting product regulation mechanism; it alters the chain length of its products depending on the cofactor present. The protein yields C10-GPP in the presence of Co2+ or Mn2+, whereas it produces the longer C15-FPP in the presence of Mg2+ [65]. That allows beetles to supply precursors for two terpene pathways, one for monoterpene metabolism (synthesis of chemical defenses) and one for sesquiterpene metabolism (JH formation), using only a single enzyme. The production of DMAPP, the allylic isomer of IPP, is catalyzed by an IPP isomerase (IPPI). Insect IPPIs require Mg2+ or Mn2+ for full catalytic activity [66, 67].

The enzymes of the MVAP are well conserved in eukaryotes; in insects all the MVAP enzymes seem to be encoded by single-copy genes, and identification of predicted amino acid sequences was possible based on sequence homology [4–6]. However biochemical characterization of purified or recombinant enzymes of the MVAP in insects is limited to HMGS [68], HMGR [69–71], IPPI [66, 67], and FPPS [60–65].

3.2. Late Steps (JH-Branch)

In the late steps of JH synthesis, conversion of FPP to farnesol (FOL) is catalyzed in D. melanogaster by a FPP phosphatase (FPPase or FPPP) [72], a member of the NagD halo alkanoic acid dehalogenase family (HAD), with orthologues in several insect species, including A. aegypti [73]. The mosquito FPPase (AaFPPase-1) is a Mg2+-dependent NagD HAD protein that efficiently hydrolyzes FPP and GPP, but not IPP [73]. Afterwards farnesol undergoes two sequential oxidation reactions that generate farnesal and farnesoic acid (FA). In mosquitoes, the first reaction is catalyzed by a short chain farnesol dehydrogenase (AaSDR-1), a member of the “classical” NADP-dependent cP2 SDR subfamily that presents broad substrate and tissue specificity [9]. Oxidation of farnesol into farnesal in mosquitoes is effected by a NAD+-dependent aldehyde dehydrogenase class 3 (AaALDH3-1) showing tissue and developmental-stage-specific splice variants [10]. Homologues of farnesol and farnesal dehydrogenases having similar activities in the CA of other insects have not yet been described.

The order of the last two biosynthetic steps, methyl esterification and epoxidation, catalyzed by a JH acid methyltransferase (JHAMT) and an epoxidase (EPOX), differs between insect species [2, 74]. In all insect species studied, recombinant JHAMTs were able to methylate JH III acid (JHA) and FA at similar rates [7, 75–79]. Homology modeling and docking simulations confirmed that JHAMT is a promiscuous enzyme capable of methylating FA and JHA [74]. In contrast, epoxidases have narrow substrate specificity; while the EPOX from the cockroach Diploptera punctata efficiently epoxidizes MF and is unable to process FA [80], Bombyx mori EPOX exhibits at least 18-fold higher activity for FA than MF [81]. Therefore, the order of the methylation/epoxidation reactions may be primarily imposed by the epoxidase’s substrate specificity [74]. In Lepidoptera, epoxidase has higher affinity than JHAMT for FA, so epoxidation precedes methylation, while in many other insects there is no epoxidation of FA but esterification of FA to form MF, followed by epoxidation to JH III.

The late steps of JH biosynthesis were generally considered to be JH-specific [2] and the identification of these enzymes was hindered by the small size of the CA gland that made their isolation and biochemical characterization difficult. All the enzymes have now been characterized in insects using molecular approaches that included EST sequencing [4, 80], mRNA differential display [7], or homology to orthologue enzymes [10, 72]. Identification of the three enzymes involved in the conversion of FPP to farnesoic acid in mosquitoes has proven that the 3 proteins are encoded by families of paralogue genes with broad substrate specificity and expression in a wide number of tissues [9, 10, 78, 82]. This is not surprising since generation of farnesol by FPPase is important beyond the CA. Farnesol and farnesal homoeostasis are vital for cells in all insect tissues, and farnesol plays important roles in the regulation of a wide variety of cell functions, including proliferation and apoptosis [83–85], while posttranslational modifications by attachment of a farnesyl group to C-terminal cysteines of target proteins by farnesyl-transferases are essential for signal transduction and vesicular transport [86]. The presence of AaFPPase, AaSDR, and AaALDH3 isozymes with several isoforms capable of catalyzing each of the 3 enzymatic reactions in mosquitoes might have facilitated the evolution of more efficient substrate specificities, as well as a better tissue and developmental regulation. On the other hand, caution needs to be applied when trying to identify orthologues of these enzymes in other insect species, as not always the closest orthologue might play the same role in the CA.

On the contrary, the last two enzymes of the pathway (JHAMT and EPOX) are encoded by single genes in most insect species and are expressed predominantly in the CA [6, 7]. It is also noteworthy that EPOX genes appear to be insect-specific and have not been found in other arthropods. EPOX genes may be an evolutionary innovation that occurred in ancestral insects for the epoxidation of MF to JH [87].

3.3. Enzymatic Activities

The development of simple methods for detailed analysis of enzymatic activities derived from insect CA is critical. Fluorescence approaches are simplifying the study of the ability of CA extracts and recombinant enzymes to metabolize MVAP and JH-branch intermediates in vitro [11, 12, 73]. Eight selected enzymes have been evaluated using mosquito CA homogenates [12]. HMGS and JHAMT have the highest activities (in the nanomolar range), while the activities of additional six enzymes are in the femtomolar range (MK, PMK, FPPS, FPPase, farnesol dehydrogenase, and farnesal dehydrogenase).

4. Regulation of CA Activity

4.1. Mechanisms of Allatoregulatory Activity

Regulatory signals control the CA at least at three different levels [88, 89]. (1) Cytological/developmental responses are the gross morphological, microscopic, or enzymatic changes that determine the overall physiological status of the glands and their maximal potential output, for example, changes in cell volume and cell number which normally proceed in conjunction with developmental changes, such as the transition to adult [90]. (2) Constitutive/long-term responses, such as variations in enzyme levels during cycles of CA activity, are measured on a time scale of several hours to days. Examples of constitutive responses are the acquisition and loss of sensitivity to allatoregulatory peptides by the CA in D. punctata [91] and A. aegypti [92]. (3) Dynamic/short-term responses are measured on a time scale of minutes or hours and can be measured readily in vitro, such as the inhibition of JH synthesis by allatostatins or the stimulation of JH synthesis by allatotropin. These responses are usually reversible upon removal of the stimulus [93].

4.2. Nutritional Regulation of JH Synthesis and the Brain

The correct allocation of nutrients between competing needs such as reproduction, growth, maturation, or flight is a vital component of an insect’s life-history strategy [94, 95]. Juvenile hormone has been described as part of a transduction system that assesses nutritional information and regulates reproduction in mosquitoes [96]. The nutrition-dependent development of the ovaries is an excellent physiological framework to understand the dynamic changes in JH biosynthesis during the gonotrophic cycle of female mosquitoes [12].

Three sources of nutrients provide energy and building blocks for the three distinct phases of ovarian development in A. aegypti. Preimaginal reserves are partially consumed during previtellogenesis (PVG); nectar-feeding adds reserves during the ovarian resting stage (ORS); and later a blood meal triggers vitellogenesis (VG) [96–101]. JH synthesis and ovarian previtellogenic maturation are activated in newly eclosed A. aegypti adult females only if teneral nutritional reserves are elevated [102]. Later, after previtellogenic maturation has been completed, JH mediates reproductive trade-offs in resting stage mosquitoes in response to nutrition [103]. Adult females A. aegypti show dynamic changes in JH biosynthesis, and regulation of the CA activity is quite different during previtellogenesis, the ovarian resting stage, and the vitellogenesis period [12] (Figure 2).

Figure 2: JH biosynthesis rates and ovarian development in female mosquitoes. Top panel: representative images of the progression of ovary development from emergence to 24 h after blood feeding. The inset in 96 h shows the lipid content of follicles from females fed 3% sugar (top) and 20% sugar (bottom). Colors for the panels match the colors for the nutrition-dependent physiological states of the CA shown in the panel below. Bottom panel: JH biosynthesis by CA dissected from pupa, sugar-fed, and blood-fed adult females. Hours represent times before (pupa) and after adult emergence (sugar-fed), or after blood feeding (BF). -axis: JH biosynthesis expressed as fmol/h. Bars represent the means ± SEM of three independent replicates of three groups of 3 CA. Colors represent the four distinct CA physiological phases identified: inactive or low activity CA (blue), active CA (black), modulated CA (green), and suppressed CA (red), from [12].

Comprehensive studies of transcripts, enzyme activities, and metabolites delimited four distinct nutrition-dependent CA physiological conditions that we named as follows: inactive, active, modulated, and suppressed CA (Figure 2) [12]. The molecular basis for JH synthesis regulation, as well as the role of brain factors or other endocrine regulators, might change during these 4 phases. We have previously described that transcript levels for most of the JH biosynthetic enzymes are very low in early pupae [6]; consequently JH synthesis rates were undetectable (below 0.5 fmol/h) in pupae 24 and 12 h before adult eclosion. Subsequently, in the last 6–8 h before adult emergence transcript levels for the biosynthetic enzymes commence to rise, the pupal CA becomes “competent” and starts to synthesize JH [6]. Although the CA of the newly emerged female is fully competent, for the next 10-11 h it synthesizes relatively low levels of JH (10 fmol/h) [12]. Decapitation during these first 12 h of imaginal life prevents increases of JH synthesis, suggesting that the brain plays a key role sensing the nutritional status and stimulating CA activity [104]. Only when preimaginal reserves are sufficient will the brain command the CA to synthesize enough JH to activate reproductive maturation [102].

Recent detailed studies in sugar-fed females revealed a previously undetected peak of maximum JH synthesis 12 h after adult emergence (Figure 2) [12]. This sharp increase in JH synthesis conveys information about teneral nutritional reserves and provides a signal to proceed with the previtellogenic maturation of the ovaries. The process of “activation” of CA is very fast and short lasting; JH synthesis increases from 10 fmol/h to almost 100 fmol/h in 2 h and decreases to less than 40 fmol/h in the next 2 h, remaining at this relatively high and constant rate until 24 h after emergence. Well-nourished females would activate the CA, increase JH synthesis levels, and complete the previtellogenic development by 48–60 h after emergence even if raised on water [104, 105].

If mosquitoes are nutritionally stressed, by 48–72 h JH synthesis is significantly reduced. This period represents the beginning of the ORS and female mosquitoes often ingest sugar meals to supplement their partially depleted preimaginal reserves. During the ORS, if nutrients are scarce, the brain directs the CA to “adjust” to the new adult nutritional condition; in mosquitoes fed a restricted diet such as 3% sugar, JH synthesis decreases to a low 12 fmol/h, triggering the resorption of ovarian follicles [95]. Decapitation during this ORS precludes this nutritional adjustment and causes significant increases in JH synthesis, emphasizing the critical role of the brain in CA nutritional modulation [104]. Finally, at 24 h after blood feeding there is an “active” suppression of JH synthesis that is critical for the completion of the vitellogenic development of the first batch of eggs and the triggering of the previtellogenic development of follicles for the second gonotrophic cycle (Figure 2) [12].

A coordinated expression of most JH biosynthetic enzymes has been previously described in mosquitoes and silkworms [6, 100, 101]. Increases or decreases in transcript levels for all the enzymes are generally concurrent with increases or decreases in JH synthesis [5, 6, 12], suggesting that transcriptional changes are at least partially responsible for the dynamic changes of JH biosynthesis. Most studies on JH synthesis have been performed using corpora allata-corpora cardiaca complexes (CA-CC). The 2 glands are very small and are intimately connected, so separating them is challenging. The synthesis of JH occurs exclusively in the CA; expression of the JH biosynthetic enzymes has been detected in the CA, but not in the CC of B. mori [106], and expression of the last 2 enzymes is also much higher in CA than CC in A. aegypti [107]. A potential role of the CC on CA regulation has been proposed in B. mori [108, 109]; separation of the CA from the CC often results in increases of JH synthesis in vitro in A. aegypti [93].

5. Allatoregulators

There are factors that can stimulate (allatotropins) or inhibit (allatostatins) CA activity [2]. In different insect species and at different stages of development, these regulatory factors may include three types of inhibitory allatostatins (AST), at least one type of stimulatory allatotropin (AT), insulin, and perhaps additional neuropeptides [110]. These factors were reviewed in detail in several recent articles [1, 2, 110–112].

5.1. Allatostatins and Allatotropins

Three families of allatostatins have been identified in insects: cockroach allatostatins (YXFGL-amide or type-A), cricket allatostatins (W2W9 or type-B), and Manduca allatostatins (PISCF or type-C) [111, 113, 114]. Each of the three structurally unrelated types of allatostatins (A, B, and C) is associated with a unique G-Protein-Coupled Receptor (GPCR) family that includes vertebrate orthologs. The AST-A receptors are related to the vertebrate galanin receptors [115], the AST-B receptors to the bombesin receptors [116], and the AST-C receptors show similarity to the somatostatin/opioid receptors [117, 118]. The AT receptor is also a GPCR and shows homology to the vertebrate orexin/hypocretin receptors [107, 108, 119, 120]. Stimulatory and inhibitory effects of brain factors have been described in mosquitoes [93, 104, 121]. Allatostatin-C and allatotropin are present in the brain of A. aegypti, [122]; they both modulate JH synthesis in vitro [92, 123] and their receptors are expressed in the CA-CC complex [107, 118]; however, their exact roles in vivo and mechanisms of action still need to be elucidated.

5.2. The Insulin/TOR Signaling Network

The insulin/TOR signaling network is evolutionarily conserved in most eukaryotes and plays a central role in the transduction of nutritional signals that regulate cell growth and metabolism [124, 125]. There are several reports suggesting that the insulin pathway modulates JH synthesis in insects. In D. melanogaster, specific silencing of the insulin receptor (InR) in the CA completely suppresses HMG-CoA reductase expression and renders a JH-deficient phenotype [126]. In addition, D. melanogaster InR mutants have reduced JH synthesis [127]. In Culex pipiens, the ability to enter into overwintering diapause is regulated by JH [128], and suppression of allatotropin simulates reproductive diapause [121]. In C. pipiens, silencing the InR or the downstream FOXO protein (forkhead transcription factor) by RNAi leads to a diapause phenotype [128]. The insulin/TOR pathway has also been suggested as a link between nutritional signals and JH synthesis regulation in the CA of the cockroach Blattella germanica [129, 130], and FOXO knockdown using systemic RNAi in vivo in starved females elicited an increase of JH biosynthesis [131].

The A. aegypti genome encodes eight insulin-like peptides (ILPs), with three of them (ILP1, ILP3, and ILP8) specifically expressed in brains of adult females [132]. ILP3 binds the A. aegypti insulin receptor (InR) with high affinity and has been described as a critical regulator of egg production [133]. Transcript levels for several A. aegypti ILPs show age-dependent and diet-dependent changes in female mosquitoes [134]. Mosquito ILPs action appears to be mediated by the tyrosine kinase activity of the mosquito insulin receptor and a signaling network involving phosphatidylinositol 3-kinase [135]. Selective activators and inhibitors of insulin signaling cascades had strong effects on insulin-regulated physiological processes in mosquitoes [135]; for example, knockdown of the A. aegypti phosphatase and tensin homolog (AaegPTEN) affects insulin signaling [136].

Application of bovine insulin on the mosquito CA-CC incubated in vitro caused a strong and fast stimulation on JH synthesis [19]. Little is known on exactly how insulin/TOR signaling affects the activity of the CA. Systemic depletion of TOR by RNAi and administration of the TOR modulator rapamycin had inhibitory effects on JH synthesis in mosquitoes, with both treatments causing reductions in JH biosynthetic enzyme transcript levels [19]. In A. aegypti, starvation decreases JH synthesis via a decrease in insulin signaling in the CA (Figure 3). Starvation-induced upregulation of the insulin receptor, increased CA insulin sensitivity and “primed” the gland to respond rapidly to increases in insulin levels. During this response to starvation, the synthetic potential of the CA remained unaffected, and the gland rapidly and efficiently responded to insulin stimulation by increasing JH synthesis to rates similar to those of CA from nonstarved females [23].

Figure 3: Starvation effects on insulin signaling components and JH synthesis in the CA of mosquitoes. This scheme summarizes starvation-related changes of insulin/TOR pathway components and JH synthesis. Molecules in red color are downregulated (↓), while those in green are upregulated (↑). Phosphoinositide 3-kinase (PI3K) and TOR are involved in the transduction of insulin signaling in the CA [13]. A starvation-dependent decrease of insulin results in an increase of FOXO signaling that promotes activation of transcription of insulin receptor (INSr) and 4E-binding protein (4EBP). Transcripts levels for FOXO increase and mRNAs for JHAMT and TOR decrease. JH synthesis decreases, while increases of 4EBP inhibit translation and increases of INSr enhance insulin sensitivity, from [23].

5.3. Additional Allatoregulatory Factors

Several additional factors have been proposed to be involved in the regulation of JH biosynthesis by the CA, including biogenic amines, 20-hydroxyecdysone (20E), ecdysis triggering hormone (ETH), and short neuropeptide F (sNPF) [2]. The steroid hormone 20E controls molting, metamorphosis, and oogenesis in insects [137–139]. 20E modulates JH synthesis in Bombyx mori larvae [140, 141], possibly by means of a direct control on the expression of some of the JH biosynthetic enzymes [109].

ETH is a small C-terminally amidated peptide, known as a major regulator of ecdysis in insects [142, 143]. Its role in inducing a stereotypical ecdysis behavioral sequence resulting in molts is well characterized [144]. ETH is synthesized and secreted into the hemolymph by specialized endocrine cells called Inka cells [142]. In A. aegypti, Inka cells are located along branch points of major epitracheal trunks [145]. The A. aegypti ETH gene encodes two isoforms of the 17 amino acid peptides, ETH1 (AeETH1) and ETH2 (AeETH2) [145]. Both of these peptides induce a receptor-mediated signaling cascade in CNS neurons that result in activation of motor programs allowing shedding of the old cuticle [142]. Yamanaka and collaborators reported very high expression of the ETH receptor in the CA of B. mori leading them to suggest that ETH might have a role in regulation of JH synthesis [108]. Preliminary results indicate a stimulatory effect of ETH on JH synthesis in A. aegypti during the maturation process of the CA in the last six hours before adult emergence, a time when genes encoding JH biosynthetic enzymes become transcriptionally active and the CA starts synthesizing basal levels of JH III [146].

The short neuropeptide F (sNPF), among other functions, modulates feeding, metabolism, reproduction, and stress responses in insects [147]. sNPF has been reported as an allatoregulatory peptide in B. mori; in the silk moth, the AT receptor is not expressed in the CA, but rather in the corpora cardiaca (CC), specifically in a group of 4 cells that express the sNPF [108]. According to the model proposed for Bombyx, AT inhibits the release of sNPF, and this peptide inhibits JH synthesis; so AT exerts an indirect allatotropic effect by “derepression.” This model has not been tested in mosquitoes or additional insect species.

In mosquitoes, the role of each of these endocrine regulators might be limited to particular periods of CA activity. Developmental modulators such as ETH might play important roles during pupal maturation of the CA; insulin and/or allatotropin may well be the brain activators acting on the CA of the newly emerged female, while allatostatin-C and insulin could play a role in the nutritional modulation of JH synthesis during the “state of arrest,” as well as during the suppression of JH synthesis after a blood meal. In the CC-CA of mosquitoes, the expression of the following receptors has been detected: ETH A and B, ecdysone A and B, insulin, ultraspiracle A and B, allatotropin, AST-C A and B, and the short neuropeptide F. It is possible that signals from all these modulators are integrated in the CA, which suggests that the regulation of JH synthesis is extremely complex (Figure 4).

Figure 4: Effect of modulators on JH biosynthesis in female mosquitoes. Schematic representation of some of the tissues and molecules involved in JH biosynthesis regulation in mosquitoes. PG: prothoracic gland. OV: ovaries. CC: corpora cardiaca. CA: corpora allata. ETH: ecdysis triggering hormone. AST-C: allatostatin-C. AT: allatotropin. INS: insulin. 20E: 20 hydroxyecdysone. R: receptor. JH: juvenile hormone. Green arrow: stimulation. Red arrow: inhibition. Black arrow: modulation.

6. An Integrated View of Flux Control of JH Synthesis Rate

6.1. Flux Control

JH synthesis is controlled by the rate of flux of isoprenoids, which is the outcome of a complex interplay of changes in precursor pools, enzyme levels, and external modulators such as nutrients and allatoregulatory factors [6, 12, 148, 149] (Figure 5). Discussion of the “control” or “regulation” of biosynthetic pathways normally focuses on the question of which individual enzymes are controlling the flux in a pathway [150, 151]. Flux is a systemic property, and questions of its control cannot be answered by looking at the different enzymatic steps in isolation. To understand how regulators modify JH synthesis, it is important to know their effect on the changes in the levels of all enzymes and precursor pool sizes.

Figure 5: A schematic representation of a model for the control of the flux of precursors in the JH biosynthetic pathway. Precursor pools (S2, S3, etc.) are represented by circles and connected by arrows (MVA: mevalonic acid, 5P-MVA: mevalonate 5-phosphate). E: enzymes are followed by a number that refers to the position in the pathway (E3 = MK: mevalonate kinase). Regulatory factors might be affecting both precursor pool sizes and enzymatic activities (e.g., AST-C: allatostatin-C). JH: juvenile hormone, from [6].

The JH synthetic pathway involves 13 discrete enzymatic steps organized in an obligatory sequence. Each product represents the substrate for the next “downstream” enzyme. Enzymes are connected by metabolite pools that are common to them; for example, FOL is the product of the FPPase activity and the substrate for farnesol dehydrogenase. The pools are in fact the links in the system interactions; therefore, pool concentrations and fluxes (which are flows into and out of pools) are critical variables in JH regulation. The system’s “sensitivity” to changes in the size of a precursor pool indicates the control importance of this enzymatic step in the final flux and can be experimentally tested. Although control of fluxes tends to be distributed among all enzymes in a pathway rather than confined to a single rate-limiting enzyme, the extent of control can differ widely between enzymes in a pathway [150]. It has been postulated that, in a synthetic pathway containing numerous enzymes, almost all the enzymes will appear to be “in excess,” in the sense that individual quantities or activities can be considerably reduced without appreciable effect on the flux [150]. Stimulation with exogenous precursors has been reported for the CA of many insect species, and it seems that having an excess of enzymes is common in most insects studied [6, 152–155]. In the CA of the cockroach Diploptera punctata, HMGS and HMGR activities are not always closely linked to the rate of spontaneous JH synthesis [154, 156]. Sutherland and Feyereisen [157] showed in D. punctata that inhibiting the HMGR activity by a third has a moderate inhibition of JH synthesis (less than 15%), indicating that this enzyme is in excess and has a low control coefficient on JH synthesis. Rate limiting bottlenecks have been proposed at single specific steps in both the MVAP and JH-branch in the CA of different insects, including upstream of the acetyl-CoA pool [157] as well as by rate limiting blockages at different enzymatic steps in the pathway, including the activities of HMGR [158, 159], farnesol dehydrogenase [9], farnesal dehydrogenase [10], or JHAMT [7, 77]. In contrast recent studies suggest that there are multiple regulatory points in the pathway and they might change in different physiological stages [12].

Branch point regulation is an important mechanism controlling carbon flow in the MVAP; the FPP produced by the MVAP can be shunted to many metabolic branches for the synthesis of critical molecules such as ubiquinone, dolichol, or prenylated proteins [59]. Remarkably, when the CA is very active, MVAP intermediate pools are completely depleted, implying that most MVAP precursors are channeled into the JH-branch. These results suggest that, during the peak of synthesis, the activity of the enzymes on the JH-branch is controlling the flux in the synthesis of JH, indicating that although CA cells are using the MVAP to synthesize additional metabolites that are important for various biological processes, when necessary, the production of JH supersedes the trafficking of FPP into other branches of the MVAP [12].

Compartmentalization of the enzymatic steps might add an additional level of complexity. Studies in plants have emphasized the importance of compartmentalization in the control of terpene biosynthesis [160], challenging the traditional view of isoprenoid metabolism occurring in a homogeneous environment with intermediates mixing freely and accessible to successive or competing enzymes. Experiments performed by Sutherland and Feyereisen [157] provided strong evidence that D. punctata CA glands inhibited with allatostatin-A (AST-A) were prevented from using glucose or amino acids to synthesize JH but free to utilize acetate; that is, AST-A was inhibiting steps in the glucose or amino acid (mitochondrial) incorporation pathway but not the acetate (cytoplasmic) incorporation pathway. Results from the D. punctata-AST-A model confirm that compartmentalization of the precursor pools and enzymatic steps is important and suggest that a major target of AST-A is either the transport of citrate across the mitochondrial membrane and/or the cleavage of citrate to yield cytoplasmic acetyl-CoA [157].

Metabolic enzymes that catalyze a series of successive reactions can form complexes on membranes or cytoskeletal structures [161]. Such metabolic enzyme complexes are called “metabolons,” functioning as metabolic channels that facilitate metabolite flux to committed end products [162]. Metabolons can “move” metabolites more efficiently through the pathway and limit the availability of potential common metabolite intermediates to other branches of the network [163]. Metabolon formation normally involves specific interactions between several “soluble” enzymes that might be anchored to a membrane either by membrane-bound structural proteins that serve as “nucleation” sites for metabolon formation or by membrane-bound proteins; AaADLH3 or epoxidase could serve that role in the CA of mosquitoes. In vertebrates, farnesal dehydrogenase closely interacts with farnesol dehydrogenase, forming a complex called “alcohol : NAD+ oxidoreductase” (FAO), responsible for the sequential oxidation of fatty alcohol to fatty acids [164, 165]. A similar complex including the two oxidoreductases, the JHAMT and epoxidase, might be present in the CA of mosquitoes, channeling precursors efficiently in the JH pathway.

In vitro experiments have shown that several intermediates in the pathway (e.g., mevalonate, farnesol, farnesal, and FA) are incorporated into the CA and stimulate JH synthesis [6, 25, 155]. It is puzzling that the CA of a newly emerged mosquito female that has a very large FA pool but limited JH synthesis is strongly stimulated by exogenous supply of FA [6, 12]. These results suggest differences in the channeling of “endogenous” and “exogenous” FA derived pools. In addition, there are examples of a reversal of the flux in the JH synthesis pathway, such as a reductase activity that converts FAL back into FOL in the CA of mosquitoes [10]. In the CA, some MVAP precursor pools might be controlled by feedback regulation imposed by metabolites such as FPP operating in the downstream portions of the pathway, in a similar mode to the negative feedback of late MVAP precursors (GPP, FPP) on the activity of mevalonate kinase described for terpene homeostasis in mammals [166].

What do integrated studies of CA transcripts, enzyme activities, and metabolites tell us about the coordination of MVAP and JH-branch activities? A comprehensive analysis of the JH biosynthetic pathway has been done in B. mori [5, 106], showing that transcripts levels for the 8 enzymes of the MVAP and JHAMT are expressed in a highly coordinated manner during the 4th and 5th instar larvae as well as in pupae and adults. There is also a coordinated expression of the 13 JH biosynthetic enzymes in pupae and adults of female mosquito [6, 12]. The mosquito studies suggest that both pathways (MVAP and JH-branch) are transcriptionally coregulated as a single unit, and catalytic activities for the enzymes of the MVAP and JH-branch also change in a coordinated fashion in the “active” and “inactive” CA [12] (Figure 6). State-of-the-art metabolic studies were implemented for the first time to measure changes in all JH precursor metabolic pools in the CA of insects [12]. Unbiased Principal Component Analyses (PCA) showed that global fluctuations in the intermediate pool sizes in the MVAP and JH-branch are not functioning as a unit but behave inversely [12]. PCA of the metabolic pools changes indicated that, in reproductive female mosquitoes, there are at least 4 developmental switches that alter JH synthesis by modulating the flux at distinctive points in both pathways (Figure 7).

Figure 6: Heat map representation of changes in JH biosynthetic enzyme mRNAs and activities in CA extracts. (a) Changes in mRNAs encoding JH biosynthetic enzymes. (b) Changes in activities of JH biosynthetic enzymes in CA extracts. Top: physiological stages are described as hours relative to adult emergence (0 h) or blood feeding (BF). Right side: enzyme names abbreviations: Acetoacetyl-CoA thiolase: thiolase; HMG-CoA synthase: HMGS; HMG-CoA reductase: HMGR; mevalonate kinase: MK; phosphomevalonate kinase: PMK; diphosphomevalonate decarboxylase: PPM-Dec; IPP isomerase: IPPI; FPP synthase: FPPS; farnesyl pyrophosphatase: FPPase; farnesol dehydrogenase: FOL-SDR; farnesal dehydrogenase: FALDH; juvenile hormone acid methyltransferase: JHAMT; and methyl farnesoate epoxidase: EPOX. Colors from white to red represent increases of transcript levels or enzymatic activities as percentages of the maximum value, from [12].

Figure 7: Schematic representation of the distinct four CA physiological conditions in reproductive female mosquitoes. The four CA phases and corresponding stages are as follows: inactive (early pupae), active (12–24 h sugar-fed females), modulated (48–96 h sugar-fed females), and suppressed (24 h blood-fed females). JH synthesis: the color and direction of the arrows reflect the following: low levels (arrows down and red), high levels (arrows up and black), or variable levels (arrow up and down). Changes in transcripts, activities, and metabolites are as follows: the direction of the arrows reflects the trend of the changes (increases: up and decreases: down); the size of the arrow reflects the magnitude of the changes, limiting factor: hypothetical critical factor limiting CA activity, from [12].

Further studies will be necessary to discover what enzymes restrict the flux into JH III at specific physiological conditions.

7. Challenges and Future Directions

JH has long been the focus of intensive research intended to exploit its properties for the purpose of generating novel pest control products. Earlier research on JH biosynthesis was performed mainly on three insect models: cockroaches, locusts, and moths. These insects offered several advantages for JH synthesis studies, such as the size of the CA, the relatively high levels of JH synthesized, and the easiness of rearing them in the laboratory. Cockroaches, in particular D. punctata, have been a favorite model because of many positive biological aspects, among them a clear correlation between cycles of JH synthesis and oocyte growth and vitellogenesis [167]. The moth M. sexta also provided an excellent endocrine system model amenable to the study of JH synthesis, in particular at the biochemical level, but did not offer the genetics necessary to further test many of the hypotheses generated by biochemical and physiological studies. The potential for genetic manipulation has made Drosophila the leader in the search for molecular mechanisms of action, but with the drawback of a lack of well-defined JH homologues and roles for JH biological activities. With the advent of genomic approaches, studies on other insect species such as Tribolium and A. aegypti are again contributing critical new insights into JH biosynthesis.

To answer the questions that remain unanswered, we need to identify some of the next challenges and future directions on JH synthesis research.(1)Although the general features of JH biosynthesis seem to be conserved in most insects, there is clearly diversity in aspects such as the presence of particular JH homologues, the order of the final enzymatic steps, and the role of allatoregulators; therefore JH biosynthesis studies need to be extended beyond the classic model insects.(2)The identification of all the genes encoding JH biosynthetic enzymes has allowed the completion of comprehensive transcriptional studies, as well as the expression and characterization of recombinant enzymes. New methods are currently facilitating the analysis of JH biosynthesis rates, enzymatic activities, and metabolite pool sizes in the CA. In the future, we should improve our understanding of the occurrence of different flux directionalities, feedback loops, and pathway branching points in the JH biosynthesis pathway.(3)More research on compartmentalization of JH synthesis is necessary, as well as a better understanding of signaling pathways in the CA, including calcium signaling pathways and the interactions among the insulin and TOR pathways.(4)The list of putative JH modulators continues to increase, and new concepts in allatoregulator-modulation of JH synthesis under different physiological frameworks are emerging.(5)The utilization of new statistical approaches, theoretical models, and system biology approaches should continue to simplify the interpretation of JH synthesis rates changes.

In summary, integrative approaches using CA metabolomics, genomics, and proteomics are promising tactics to identify regulatory points in the flux of precursors in the JH synthesis pathway and unveil the molecular mysteries of a complex metabolic system such as the synthesis of juvenile hormone in the corpora allata of insects.

Conflict of Interests

The author declares that there is no conflict of interests regarding the publication of this paper.


The author thanks Dr. Mark Clifton, Dr. Martin Edwards, Dr. Crisalejandra Rivera Perez, and Mr. Pratik Nyati for critical reading of the paper. This work was supported by NIH Grant no. AI 45545 to Fernando G. Noriega.

The Quorum-Sensing Molecule Farnesol Is a Modulator of Drug Efflux Mediated by ABC Multidrug Transporters and Synergizes with Drugs in Candida albicans


Overexpression of the CaCDR1-encoded multidrug efflux pump protein CaCdr1p (Candida drug resistance protein 1), belonging to the ATP binding cassette (ABC) superfamily of transporters, is one of the most prominent contributors of multidrug resistance (MDR) in Candida albicans. Thus, blocking or modulating the function of the drug efflux pumps represents an attractive approach in combating MDR. In the present study, we provide first evidence that the quorum-sensing molecule farnesol (FAR) is a specific modulator of efflux mediated by ABC multidrug transporters, such as CaCdr1p and CaCdr2p of C. albicans and ScPdr5p of Saccharomyces cerevisiae. Interestingly, FAR did not modulate the efflux mediated by the multidrug extrusion pump protein CaMdr1p, belonging to the major facilitator superfamily (MFS). Kinetic data revealed that FAR competitively inhibited rhodamine 6G efflux in CaCdr1p-overexpressing cells, with a simultaneous increase in an apparent Km without affecting the Vmax values and the ATPase activity. We also observed that when used in combination, FAR at a nontoxic concentration synergized with the drugs at their respective nonlethal concentrations, as was evident from their <0.5 fractional inhibitory concentration index (FICI) values and from the drop of 14- to 64-fold in the MIC80 values in the wild-type strain and in azole-resistant clinical isolates of C. albicans. Our biochemical experiments revealed that the synergistic interaction of FAR with the drugs led to reactive oxygen species accumulation, which triggered early apoptosis, and that both could be partly reversed by the addition of an antioxidant. Collectively, FAR modulates drug extrusion mediated exclusively by ABC proteins and is synergistic to fluconazole (FLC), ketoconazole (KTC), miconazole (MCZ), and amphotericin (AMB).

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Infections caused by the dimorphic opportunistic pathogen Candida albicans are treated by antifungal agents. Widespread and prolonged usage of antifungals, in recent years, has led to the emergence of strains of Candida which display multidrug resistance (MDR) (15, 16, 19). Among the various mechanisms used by the fungus to gain resistance toward antifungal therapy, enhanced drug export represents an important strategy. Most of the clinically drug-resistant isolates of C. albicans are shown to overexpress genes encoding CaCDR1, CaCDR2, or CaMDR1 drug efflux pump proteins. CaCDR1 and CaCDR2 belong to ATP binding cassette (ABC) transporters, which use energy driven from ATP hydrolysis to transport drugs outside the cells, while CaMDR1, a major facilitator superfamily (MFS) protein, utilizes proton gradient for drug extrusion (20, 23). Notably, major multidrug transporters of Candida that belong to different superfamilies of proteins are functionally identical in expelling drugs but differ mechanistically in achieving drug expulsion. Among various strategies employed to combat MDR, blocking or modulating the function of the drug efflux pump proteins represents an attractive approach (35).

MDR in cancer cells is an obstacle to effective chemotherapy. ABC transporters, including ABCB1, ABCC1, and ABCG2, play an important role in the development of frequently encountered MDR in cancer cells (29). Here again, among different approaches employed to overcome MDR, inhibition of the drug extrusion pump activity represents an attractive approach (29, 36). Many clinically relevant anticancer drugs, such as Vinca alkaloids (vinblastine and vincristine), anthracyclines (doxorubicin and daunorubicin), taxenes (paclitaxel and docetaxel), epipodophylltoxins (etoposide and teniposide), camptothecins (topotecan), and anthracenes, are identified as modulators of human ABC transporters which offer great hope in successful cancer chemotherapy (36). In comparison, modulators of MDR pump proteins in pathogenic yeasts are only beginning to be characterized. There are already examples of compounds, such as enniatins, milbemycins, synthetic d-octapeptides, isonitrile, and unnarmicins, which modulate drug efflux by inhibiting the fungal multidrug transporters (11, 35). We have earlier shown that disulfiram, an antabuse, acts as a modulator of CaCdr1p by inhibiting oligomycin-sensitive ATP hydrolysis and affecting drug binding sites in CaCdr1p (33). Recently, polyphenol curcumin (CUR) has also been shown to be a specific modulator of rhodamine 6G (R6G) efflux mediated by CaCdr1p, CaCdr2p, and ScPdr5p (27). CUR competitively inhibited R6G efflux and the photolabeling of CaCdr1p by the prazosin analog [125I]iodoarylazidoprazosin without affecting ATPase activity (27).

Farnesol (FAR), a quorum-sensing molecule (QSM), is a precursor for the synthesis of sterols in C. albicans; it also blocks the morphological transition and biofilm development in Candida (10). FAR is known to be involved in triggering apoptosis in human oral squamous carcinoma cells (24). In mammalian cells, FAR interferes with calcium signaling and membrane fluidity (24). Studies on quorum sensing suggest its involvement in fungus-bacterium interactions and biofilm formation (34). Notably, FAR also induces apoptosis in a number of fungal species (4, 25). A global protein expression profiling following FAR treatment in C. albicans revealed mitochondrial degradation, reactive oxygen species (ROS) accumulation, caspase activation, and apoptosis as a cause of cell death (30). In this study, we provide evidence that FAR could also specifically modulate drug extrusion mediated by ABC transporters, such as CaCdr1p and CaCdr2p, without affecting the MFS transporter, such as CaMdr1p. It specifically modulates the efflux of substrates, such as R6G and fluconazole (FLC), whereas it has no effect on the efflux of substrates like Nile red (NR) and methotrexate (MTX). FAR at its nonlethal concentrations also synergizes with azoles and polyenes. Together, we show that FAR is a specific modulator of the efflux of drugs mediated by ABC transporter proteins, and it also displays synergism to antifungals by accumulating ROS and resulting in an early cell death.

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Materials.Rhodamine 6G (R6G), 2,4-dinitrophenol (DNP), 2-deoxy-d-glucose (DOG), oligomycin, 3-(4,5-dimethyl thiazol-2-yl)-2,5 diphenyl tetrazolium bromide (MTT), Nile red (NR), and other molecular-grade chemicals were obtained from Sigma Chemicals Co. (St. Louis, MO). [3H]fluconazole ([3H]FLC; specific activity, 19 Ci/mmol) was custom synthesized from Amersham Biosciences, United Kingdom, and [3H]methotrexate ([3H]MTX; specific activity, 8.60 Ci/mmol) was procured from Amersham Biosciences, United Kingdom. 2′,7′-Dichlorofluorescin diacetate (DCFH-DA), ascorbic acid (AA), and other molecular-grade chemicals were obtained from Sigma Chemicals Co. (St. Louis, MO). The annexin V-fluorescein isothiocyanate (FITC) apoptosis detection kit was obtained from BD Biosciences.

Yeast strains and growth media.The strains used in this study are listed in Table 1. The yeast strains were cultured in yeast extract-peptone-dextrose (YEPD) broth (BIO101; Vista, CA) or RPMI 1640 medium. For agar plates, 2.5% (wt/vol) Bacto agar (Difco, BD Biosciences, NJ) was added to the medium. All strains were stored as frozen stocks with 15% glycerol at −80°C. Before each experiment, cells were freshly revived on YEPD plates from the stock.

Table 1.

Strains used in this study

Efflux of rhodamine 6G.Efflux of R6G was determined essentially using a previously described protocol (32). Briefly, approximately 1 × 106 yeast cells from overnight-grown culture were transferred into YEPD medium and allowed to grow for 5 h. Cells were pelleted, washed twice with phosphate-buffered saline (PBS; without glucose), and resuspended as a 2% cell suspension, which corresponds to 108 cells (wt/vol) in PBS without glucose. The cells were then deenergized for 45 min in DOG (5 mM) and DNP (5 mM) in PBS (without glucose). The deenergized cells were pelleted, washed, and then resuspended as a 2% cell suspension (wt/vol) in PBS without glucose, to which R6G was added at a final concentration of 10 μM and incubated for 40 min at 30°C. The equilibrated cells with R6G were then washed and resuspended as a 2% cell suspension (wt/vol) in PBS without glucose. Samples (1-ml volumes) were withdrawn at the indicated time and centrifuged at 9,000 × g for 2 min. The supernatant was collected, and absorption was measured at 527 nm. Energy-dependent efflux (at the indicated time shown in Fig. 1) was measured after the addition of glucose (2%) to the cells resuspended in PBS (without glucose). Glucose-free controls were included in all the experiments. For competition assays, FAR (100 μM) was added to the deenergized cells 5 min before the addition of R6G and allowed to equilibrate.

Measurement of drug accumulation.The accumulation of [3H]FLC (specific activity, 19 Ci/mmol), [3H]MTX (specific activity, 8.60 Ci/mmol), and fluorescent NR was determined essentially by the methods described previously (17, 21). Briefly, cells from mid-log phase (5 × 106) were centrifuged at 3,000 × g for 3 min and resuspended in PBS as a 2% cell suspension. For accumulation studies, 100 nM FLC and 25 μM MTX were routinely used (17). FAR (100 μM) was added 5 min before the addition of drugs and allowed to equilibrate. One hundred microliters of cell suspension containing drugs or drugs with FAR were incubated at 30°C for 40 min, filtered rapidly, and washed twice with PBS (pH 7.4) on a Millipore manifold filter assembly using a 0.45-μm-pore-size cellulose nitrate filter (Millipore). The filter discs were dried and put in cocktail O, and the radioactivity was measured in a liquid scintillation counter (Beckman). The accumulation was expressed as picomoles/milligram (dry weight). Accumulation of NR was measured by flow cytometry with a FACsort flow cytometer (Becton-Dickson Immunocytometry Systems, San Jose, CA) as described previously (21).

ATPase assay.ATPase activity of the plasma membrane fractions was measured as oligomycin-sensitive release of inorganic phosphate either alone or in the presence of FAR (100 μM) as described previously (32).

Time-kill assays.C. albicans cells at a concentration of 103 CFU/ml were inoculated in RPMI 1640 medium containing either FAR or antioxidant ascorbic acid (AA) alone or a combination of both FAR and AA as detailed previously (26). At predetermined time points (0, 4, 8, 12, 16, 20, and 24 h) (at 30°C incubation; agitation, 200 rpm), a 100-μl aliquot was removed, serially diluted (10-fold) in 0.9% saline, and plated on YEPD agar plates. Colony counts were determined after incubation at 30°C for 48 h (26).

Measurement of ROS production.Endogenous amounts of ROS were measured by a fluorometric assay with 2′,7′-dichlorofluorescin diacetate (DCFH-DA) (26). Briefly, the cells were adjusted to an optical density at 600 nm (OD660) of 1 in 10 ml of PBS and centrifuged at 2,500 × g for 15 min. The cell pellet was then resuspended in PBS and treated with appropriately diluted AA for 1 h or was left untreated at room temperature. After incubation with FAR at 37°C for different time intervals as indicated, 10 μM DCFH-DA in PBS was added. The fluorescence intensities (excitation and emission of 485 and 540 nm, respectively) of the resuspended cells were measured with a spectrofluorometer (Varian; Cary Eclipse).

Analysis of apoptotic markers.Protoplasts of C. albicans were stained with propidium iodide (PI) and FITC-labeled annexin V by using the annexin V-FITC apoptosis detection kit (BD Biosciences) to assess cellular integrity and the externalization of phosphatidylserine (PS) as described earlier (26). The cells were analyzed by using a fluorescence-activated cell sorter (FACS) caliber flow cytometer (Becton Dickinson Immunocytometry Systems, San Jose, CA) using a 488-nm excitation and a 515-nm band pass filter for FITC detection and a filter >560 nm for PI detection. A total of 10,000 events were counted at the flow rate. Data analysis was performed using Cell Quest software (Becton Dickinson Immunocytometry Systems) (26).

Liquid susceptibility assay.The interaction of FAR with FLC, ketoconazole (KTC), miconazole (MCZ), and amphotericin (AMB) was evaluated by the checkerboard method recommended by the NCCLS and expressed as the sum of the fractional inhibitory concentration index (FICI) for each agent. The FIC of each agent is calculated as the MIC of this agent in combination divided by the MIC of this agent alone (26). In brief, serial double dilutions of the anticandidal compounds were prepared ranging from 0.25 to 128 μg/ml for FLC, 0.019 to 10 μg/ml for KTC, and 0.019 to 10 μg/ml for MCZ and AMB. After drug dilutions were made, a 100-μl suspension of Candida strains adjusted to 5 × 105 CFU/ml was added to each well and cultured at 30οC for 48 h in RPMI 1640 medium. Then visual reading of MICs was performed, and OD600 values were measured. The background OD value was subtracted from the OD value of each well. Each checkerboard test generates many different combinations, and by convention the FIC value of the most effective combination is used in calculating the FIC index. FICI was calculated by adding both FICs: FICI = FICA + FICB = CAcomb/MICAalone + CBcomb/MICBalone, where MICAalone and MICBalone are the MICs of drug A and B when acting alone and CAcomb and CBcomb are concentrations of drugs A and B at the isoeffective combinations, respectively. Off-scale MICs were converted to the next highest or next lowest doubling concentration. The FICI was interpreted as synergistic when it was ≤0.5, as antagonistic when >4.0, and as indifferent at any value in between (26).

MTT assay.The cytotoxic effect of FAR was determined by MTT assay (1, 2, 30). Yeast cells (104) were seeded into 96-well plates in the absence and in the presence of various concentrations of FAR (25 to 500 μM) and grown for 48 h at 30°C. MTT solution (100 μl) was added to each well and incubated for 3 to 4 h, and 200 μl of isopropanol was added to stop the reaction. The absorbance of the whole microtiter plate was measured using a microplate spectrophotometer at 570 nm with a reference wavelength of 650 nm. Cell survival (as a percentage of the control) equals the mean absorbance in the test well divided by the mean absorbance in control wells, multiplied by 100.

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FAR inhibits R6G efflux mediated by ABC transporters.In this study, we explored the modulator effect of FAR on MDR efflux proteins of C. albicans. For this, we monitored the transport of well-known efflux substrate R6G (10 μM) in the presence of FAR (100 μM) in cells where green fluorescent protein (GFP)-tagged ABC transporters, such as CaCdr1p (CDR1-GFP), CaCdr2p (CDR2-GFP), and ScPdr5p (PDR5-GFP), were stably overexpressed from a genomic PDR5 locus in the Saccharomyces cerevisiae AD1-8u mutant, which is deleted in seven ABC transporters (15). We have shown earlier that overexpression of GFP-tagged ABC proteins of C. albicans offered expression levels sufficient for the biochemical characterization of the transporters (17, 32). It should be mentioned that the functionality of the GFP-tagged version of MDR transporters remains similar to their respective untagged proteins (13, 17, 32). As depicted in Fig. 1, there was no efflux of R6G in control energized AD1-8u cells. In contrast, the AD1-8u cells overexpressing CDR1-GFP showed time- and energy-dependent efflux of R6G. This was evident from a steady increase in the extracellular concentration of R6G (Fig. 1A). We further evaluated if the observed inhibitory effects of FAR on R6G transport could be extended to CaCdr1p homologues, such as CaCdr2p and ScPdr5p, which were also expressed in similar heterologous backgrounds (15). It was observed that FAR could inhibit the efflux of R6G mediated by both the proteins (Fig. 1B and C). Of note, FAR could also modulate R6G efflux in C. albicans wild-type (WT) (SC5314) cells (Fig. 1D); however, for subsequent detailed studies, we used the heterologous host AD1-8u strain overexpressing MDR transporters.

Fig. 1.

Effect of FAR on R6G transport. Extracellular R6G concentrations in S. cerevisiae control cells (AD1-8u) and in cells overexpressing CaCdr1p (AD-CDR1) (A), CaCdr2p (AD-CDR2) (B), and ScPdr5p (AD-PDR5) (C). (D) Wild-type strain SC5314 of C. albicans. The energy-dependent R6G efflux was initiated by adding glucose (2%; indicated by an arrow) and quantified by measuring the absorbance of the supernatant at 527 nm. The values are the means ± standard deviations (indicated by error bars) from three independent experiments.

FAR is a substrate-specific modulator of ABC proteins.While FAR was able to modulate R6G efflux mediated by all the tested ABC transporter proteins, such as CaCdr1p, CaCdr2p, and ScPdr5p (Fig. 1A, B, and C), we explored if FAR could also affect other known substrates of ABC transporters, such as FLC, MTX, and NR. As depicted in Fig. 2A, there was no change in the accumulation of FLC in AD1-8u control host cells, while cells overexpressing CaCdr1p showed reduced accumulation of FLC (enhanced efflux). Interestingly, the presence of FAR (100 μM) blocked the transport of FLC (∼80%) mediated by CaCdr1p-expressing cells, which was evident by its increased accumulation (decreased efflux). As depicted in Fig. 2B and C, the S. cerevisiae cells overexpressing CaCdr1p showed decreased accumulation of MTX and NR (increased efflux), respectively; however, unlike in the case of FLC, the presence of FAR (100 μM) had no effect on MTX or NR transport.

Fig. 2.

FLC, MTX, and NR transport in CaCdr1p-overexpressing S. cerevisiae cells. (A) [3H]FLC accumulation in S. cerevisiae control cells (AD1-8u) and in cells overexpressing CaCdr1p (AD-CDR1). Cells were incubated with either [3H]FLC (100 nM; specific activity, 19 Ci/mmol) or [3H]FLC and FAR (100 μM). The accumulated [3H]FLC was measured 40 min after the addition of glucose (2%). The values indicated by the bars represent the means ± standard deviations (indicated by error bars) from three independent experiments. (B) [3H]MTX accumulation in S. cerevisiae control cells (AD1-8u) and in cells overexpressing CaCdr1p (AD-CDR1). Cells were incubated with either [3H]MTX (25 μM; specific activity, 8.60 Ci/mmol) or [3H]MTX and FAR (100 μM). The accumulated [3H]MTX was measured 40 min after the initiation of efflux, using a liquid scintillation counter (Beckman). The values indicated by the bars represent the means ± standard deviations (indicated by error bars) from three independent experiments. (C) NR accumulation in S. cerevisiae control cells (AD1-8u) and in cells overexpressing CaCdr1p (AD-CDR1). Cells were incubated with either NR (3.5 μM) or NR and FAR (100 μM). The accumulated NR was measured 40 min after the initiation of efflux.

FAR has no effect on the transport mediated by MFS protein.We explored if FAR could also affect a multidrug transporter belonging to the MFS superfamily, and we examined the efflux mediated by CaMdr1p (an MFS transporter) expressed in similar heterologous backgrounds. For this, we monitored the transport of well-known substrates of CaMdr1p, such as FLC, MTX, or NR. As shown in Fig. 3, FAR could not block the efflux of FLC (Fig. 3A), MTX (Fig. 3B), and NR (Fig. 3C) mediated by CaMdr1p overexpressed in similar backgrounds.

Fig. 3.

FLC, MTX, and NR transport in CaMdr1p-overexpressing S. cerevisiae cells. (A) [3H]FLC accumulation in S. cerevisiae control cells (AD1-8u) and in cells overexpressing CaMdr1p (AD-CaMDR1). Cells were incubated with either [3H]FLC (100 nM; specific activity, 19 Ci/mmol) or [3H]FLC and FAR (100 μM). The accumulated [3H]FLC was measured 40 min after the addition of glucose (2%). The values indicated by the bars represent the means ± standard deviations (indicated by error bars) from three independent experiments. (B) [3H]MTX accumulation in S. cerevisiae control cells (AD1-8u) and in cells overexpressing CaMdr1p (AD-CaMDR1). Cells were incubated with either [3H]MTX (25 μM; specific activity, 8.60 Ci/mmol) or [3H]MTX and FAR (100 μM). The accumulated [3H]MTX was measured 40 min after the initiation of efflux, using a liquid scintillation counter (Beckman). The values indicated by the bars represent the means ± standard deviations (indicated by error bars) from three independent experiments. (C) NR accumulation in S. cerevisiae control cells (AD1-8u) and in cells overexpressing CaMdr1p (AD-CaMDR1). Cells were incubated with either NR (3.5 μM) or NR and FAR (100 μM). The accumulated NR was measured 40 min after the initiation of efflux.

FAR is not a substrate of MDR pumps.The fact that FAR inhibits drug transport suggests that FAR could be acting as a competing substrate for the MDR transporters. Therefore, cytotoxicity of FAR in the control (AD1-8u) and ABC/MFS transporter-expressing cells (AD-CDR1, AD-CDR2, AD-PDR5, AD-CaMDR1) was determined by using MTT assay. As shown in Fig. 4, the 50% inhibitory concentrations (IC50s) of FAR for control host strain AD1-8u, AD-CDR1, AD-CDR2, AD-PDR5, and AD-CaMDR1 were 432.44 ± 3.132, 377.99 ± 3.090, 415.66 ± 2.957, 402.87 ± 4.583, and 421.14 ± 3.355 μM, respectively. The relative resistance factor was between ∼0.874 and 1.0 (Fig. 4), indicating that the overexpression of different pump proteins could not affect IC50s of FAR. This implies that FAR could not be transported by both the control cells and the efflux pump protein-overexpressing cells. As depicted in Fig. 4, the IC50s for control (AD1-8u) cells and cells expressing the transporters (ABC or MFS) were not very different up to 400 μM FAR, ensuring that at the tested concentration of FAR, the viability of cells was not affected.

Fig. 4.

Effect of FAR on the viability of S. cerevisiae cells overexpressing MDR pumps as determined by MTT assay. (A) Percent cell survival in control cells (AD1-8u) and in cells overexpressing ABC/MFS transporters. The experiments were conducted in triplicates, and the values represent means ± standard deviations from three independent experiments. The table depicts the IC50s and the relative resistance factors for AD1-8u, AD-CDR1, AD-CDR2, AD-PDR5, and AD-CaMDR1 in the presence of FAR.

FAR competitively inhibits R6G efflux.The efflux of R6G was inhibited by FAR in a concentration-dependent manner, with an IC50 of 30 ± 5 μM (Fig. 5A). The Lineweaver-Burk plot analysis revealed that FAR competitively inhibited R6G efflux with an increase in apparent Km (6.39 to 17.68 μM) and with no change in the Vmax values (Fig. 5B). Notably, FAR at the modulator concentration did not affect the ATPase activity of CaCdr1p (Fig. 5C) and had no effect on the leakage of R6G (Fig. 1A).

Fig. 5.

Biochemical analysis of CaCdr1p in the presence of FAR. (A) For the competition assay of R6G and FAR, CaCdr1p-overexpressing S. cerevisiae cells were incubated with either R6G (10 μM) or R6G (10 μM) and FAR (10 to 100 μM). R6G efflux was monitored 40 min after the addition of glucose (2%). (B) Lineweaver-Burk plot of CaCdr1p-mediated R6G efflux in the presence of FAR 5 min after the addition of glucose (2%). The x axis (1/S) represents the various concentrations (μM) of R6G used, and the y axis (1/V) shows the rate of release of R6G in the absence (0X) and in the presence of 50 μM (5X) and 100 μM (10X) of FAR. The rate of each reaction was calculated as nanomoles of R6G released/minute/5 × 106 cells. (C) Effect of FAR on the ATPase activity of CaCdr1p. Plasma membranes from cells overexpressing CaCdr1p were incubated with and without 100 μM FAR in the ATPase buffer. The assay was performed essentially as described in Materials and Methods.

FAR is synergistic to tested drugs.FAR is known to show antifungal activity against various species of Candida (30). Here, we confirm that FAR elicited antifungal activity against the WT strain and azole-resistant clinical isolates of Candida. In addition, we also observed that FAR displays synergistic interactions with the known antifungals. A positive interaction with the drugs as demonstrated by checkerboard assays was observed. For example, in the WT strain of C. albicans, the MIC80 values of FLC, KTC, MCZ, and AMB alone were 0.5, 0.5, 0.2, and 0.31 μg/ml, respectively, which in combination with FAR was reduced to 0.00775, 0.062, 0.025, and 0.234 μg/ml, resulting in 64-, 8-, 8-, and 16-fold drops in the MIC80 values, respectively. A FICI value of less than 0.5 suggested synergism between the tested drugs and FAR (Table 2). The time-kill curves in the presence of the drugs at synergistic concentrations either alone or in combination confirmed the checkerboard results (Fig. 6).

Fig. 6.

Time-kill curves in the presence of drugs and FAR in the WT strain SC5314. FAR in combination with (i) FLC or FLC and AA, (ii) KTC or KTC and AA, (iii) MCZ or MCZ and AA, and (iv) AMB or AMB and AA. The concentrations used are FLC (0.00775 μg/ml) and FAR (11 μg/ml), KTC (0.062 μg/ml) and FAR (5.5 μg/ml), MCZ (0.025 μg/ml) and FAR (22 μg/ml), and AMB (0.078 μg/ml) and FAR (11 μg/ml). Ascorbic acid (AA) is used at a concentration of 25 mM. CNT, control having no drug.

Table 2.

Checkerboard assay of FLC, KTC, MCZ, AMB, and FAR against the WT strain of C. albicans in the absence and presence of AA

FAR at nontoxic concentrations display synergy with drugs in azole-resistant isolates.The synergistic effect of FAR in combination with FLC/KTC/MCZ/AMB could be extended against azole-resistant isolates of C. albicans, viz., Gu5 and F5. The Gu5 and F5 isolates represent AR strains derived from azole-sensitive isolates Gu4 and F2, respectively (5, 6). Both Gu5 and F5 strains display high MIC80 values, which is predominantly due to an overexpression of CaCDR1 and CaMDR1, respectively (5, 6). FAR showed synergistic interactions with the drugs in clinical isolates Gu5 and F5, as demonstrated by FICI values of less than 0.5 (Table 3). The considerable drop in the MIC80 values of the drugs (1.5- to 16-fold) in the clinical isolates points to the efficacy of FAR even in azole-resistant clinical isolates. Further, these interactions were confirmed by the time-kill curves (see Fig. S1 and S2 in the supplemental material).

Table 3.

Checkerboard assay of FLC, KTC, MCZ, AMB, and FAR against azole-resistant isolates Gu5 and F5

The synergism between FAR and drugs raises ROS levels.As depicted in Fig. 7A, the treatment of drugs such as FLC/KTC/MCZ/AMB or FAR alone at nontoxic, synergistic concentrations did not influence the levels of endogenous ROS; however, in combination with the drugs, FAR considerably augmented ROS levels. Since the combination of drugs and FAR generated ROS, we speculated whether reversal of it could affect the synergism. For this, we performed checkerboard assays in the presence of an antioxidant ascorbic acid (AA) and could indeed observe a partial reversal of synergism with a simultaneous decrease in ROS levels (Table 2 and Fig. 7A).

Fig. 7.

ROS levels in the presence of FAR/drugs and detection of apoptosis. (A) Amounts of ROS produced in WT strain SC5314 following treatment with either drugs alone or in combination with FAR or FAR and AA. The drugs (concentrations) used are FLC (0.00775 μg/ml) and FAR (11 μg/ml), KTC (0.062 μg/ml) and FAR (5.5 μg/ml), MCZ (0.025 μg/ml) and FAR (22 μg/ml), and AMB (0.078 μg/ml) and FAR (11 μg/ml). AA is used at a concentration of 25 mM. FAR-F, FAR-K, FAR-M, and FAR-A show ROS generated due to FAR alone when used at a concentration of 11 μg/ml, 5.5 μg/ml, 22 μg/ml, and 11 μg/ml, respectively. (B) Detection of apoptosis after costaining of cells with annexin V and PI in control untreated WT strain SC5314 and in cells following treatment with either FLC or AMB either alone or in combination with FAR or FAR and AA. The drugs (concentrations) used are FLC (0.00775 μg/ml), FAR (11 μg/ml), AMB (0.078 μg/ml), and AA (25 mM). The concentrations used are chosen from the checkerboard data.

Increased ROS leads to apoptosis.FAR produced by C. albicans has the ability to induce apoptosis via ROS generation and upregulation of a metacaspase, MCA1, that is involved in apoptosis or programmed cell death (PCD) (30). In addition, our previous studies of drug-drug interactions have shown that raised ROS levels due to synergy between drugs and polyphenol CUR lead to apoptosis (26). We checked if the augmented ROS due to the combination of FAR and tested drugs triggers apoptosis. To explore this, we selected two representative drugs, viz., FLC and AMB, for further analysis. As shown in Fig. 7B, there was almost no apoptotic population (∼1.5 to 2%) in cells treated with nontoxic synergistic concentrations of FAR, FLC, or AMB alone. However, there was a significant population of apoptotic cells when FAR was used in combination with FLC (18.44%) or AMB (10.44%). The treatment of cells with AA could partly reverse the percentage of apoptotic population (4.22% and 3.15% in FLC-FAR and FLC-AMB combinations, respectively).

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Among the various transporter genes identified in the C. albicans genome (7, 8), there are overwhelming clinical and experimental evidences to suggest that out of 28 ABC transporters, only two of its members, CaCdr1p and CaCdr2p, and only CaMdr1p out of a total 95 of members of MFS transporters are major determinants of MDR (18, 23). Therefore, the search for novel and potent modulators which can block and reverse the drug extrusion mediated by these efflux proteins represents an attractive strategy of anti-Candida therapy. While there are several compounds which are shown to reverse MDR mediated by Pgp (human homologue of CaCdr1p and CaCdr2p) and are at various stages of clinical trials, there are not many instances of compounds which could inhibit/reverse drug extrusion mediated by fungal multidrug transporters (12, 14, 22, 31). We have been exploring modulators/inhibitors of MDR pump proteins which could block the efflux of drugs from fungal cells. For example, we have earlier reported that purified polyphenol CUR and disulfiram, a drug used in treating alcoholism, act as modulators of CaCdr1p and thus could reverse drug efflux from Candida cells (27, 33). However, the mechanisms by which these two modulators affect the functioning of the drug efflux pump seem to differ. For example, while CUR competes for the drug binding sites without affecting the ATPase activity (27), disulfiram blocks the drug efflux by competing with the substrate binding sites as well as with a simultaneous abrogation of ATPase activity of CaCdr1p (33).

In the present study, we provide first evidence that a quorum-sensing, antifungal FAR is a modulator of drug efflux pump proteins of Candida. Thus, FAR could reverse the extrusion of specific compounds mediated exclusively by the ABC drug transporters, such as CaCdr1p, CaCdr2p, and ScPdr5p, since the efflux mediated by an MFS transporter CaMdr1p could not be inhibited by FAR. This selectivity of modulators to one type of superfamily of transporters is not uncommon (27). We had earlier observed that the modulator CUR could affect the efflux activity mediated only by the ABC drug transporters (27). If one considers the physiological concentration of FAR, which ranges between 30 and 40 μM (10), and compares it with the IC50 of the drug efflux modulation concentration (30 μM), it implies that in vivo FAR, in addition to being a QSM, could also modulate transport.

We show that FAR is not a transport substrate of ABC transporter CaCdr1p but that it can specifically modulate efflux of R6G and FLC mediated by it. This is not surprising, since it has been observed that several modulators could inhibit drug transport without being a substrate of efflux pump proteins (27, 33). Our relative resistance factor values from the cytotoxicity data (Fig. 4) reinforce the fact that the presence or absence of efflux pump proteins did not confer additional advantage toward resistance against FAR or affected the growth and viability of transporter-expressing cells. Our data that FAR is not a transport substrate but that it still competitively modulates R6G transport without affecting the ATPase activity in CaCdr1p suggests that there are either common (to R6G) or independent binding sites of FAR on CaCdr1p. However, elaborate binding studies will be required to resolve this issue.

When a nontoxic concentration of FAR was used in combination, it displayed synergy with the drugs in Candida cells. Notably, FAR could also synergize with the drugs in azole-resistant (with high MIC80 values) Candida isolates. It should be pointed out that the modulator effect of FAR over the efflux of drugs mediated by ABC multidrug transporter proteins was independent of its ability to show synergism with the drugs. For example, the synergistic action of nontoxic FAR (100 μM/22 μg ml−1) in combination with the nonlethal concentration of drugs was associated with the substantial accumulation of ROS levels (Fig. 7A), which was not the case when FAR alone at its modulator concentration was used. Our observation that FAR acts synergistically with the tested drugs suggests a possibility wherein the combination could be used to inhibit biofilm development in C. albicans. However, this warrants further studies on biofilm development at the synergistic concentrations of FAR and drugs. We had earlier shown that CUR acts as an antifungal agent against C. albicans and is synergistic to azoles (FLC) or polyenes (AMB) via generation of ROS and induction of PCD of C. albicans cells (28). Thus, various effects of CUR seem to mimic the action of FAR. However, we observed that the CUR effect was independent of FAR levels. In a mutant strain of C. albicans which was a knockout of dpp3dpp3 mutant), which encodes a phosphatase for converting farnesyl pyrophosphate to FAR, polyphenol CUR continued to inhibit cell growth, which could be reversed by the addition of an antioxidant (28). Collectively, we demonstrate that FAR is a substrate-specific modulator of efflux of drugs mediated by ABC transporter proteins and, if given in combination, is synergistic to drugs and accumulates ROS, resulting in early PCD.

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The work presented in this paper has been supported in part by grants to R.P. from the Department of Biotechnology (grant no. BT/PR11158/BRB/10/640/2008 and BT/PR13641/MED/29/175/2010) and the Department of Science & Technology (grant no. SR/SO/BB/0034/2008). M.S acknowledges the Department of Biotechnology (DBT), India, for the award of junior and senior research fellowships.

We thank Ranbaxy Laboratories Limited, India, for providing fluconazole.

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    • Received 15 March 2011.
    • Returned for modification 11 April 2011.
    • Accepted 8 July 2011.
    • Accepted manuscript posted online 18 July 2011.
  • ↵*Corresponding author. Mailing address: Membrane Biology Laboratory, School of Life Sciences, Jawaharlal Nehru University, New Delhi 110067, India. Phone: 91-11-26704509. Fax: 91-11-26741081. E-mail: rp47{at}mail.jnu.ac.in.
  • † Supplemental material for this article may be found at http://dx.doi.org/10.1128/AAC.00344-11.


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