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Evaluation of perfluorooctanoate for potential genotoxicity

Perfluorooctanoate (PFOA) is a fully fluorinated eight-carbon fatty acid analog with exceptional stability toward degradation that has been used as an industrial surfactant and has been detected in environmental and biological matrices. Exposures to PFOA in the workplace and in the environment have continuously stimulated investigations into its potential human health hazards. In this article, the results of fifteen unpublished genotoxicity assays conducted with perfluorooctanoate (as either the linear or linear/branched ammonium salt (APFO) or the linear/branched sodium salt) are reported and include: seven mutation assays (three in vitro reverse mutation assays with histidine auxotrophic strains of Salmonella typhimurium, two in vitro reverse mutation assays with the tryptophan auxotrophic Escherichia coli WP2uvr strain, one in vitro mitotic recombination (gene conversion) assay with Saccharomyces cerevisiae D4, and an in vitro Chinese hamster ovary (CHO) HGPRT forward mutation assay); seven studies to assess potential for chromosomal damage (three in vitro CHO chromosomal aberration studies, an in vitro human whole blood lymphocyte chromosomal aberration study, and three in vivo mouse micronucleus assays); and an in vitro C3H 10T1/2 cell transformation assay. Although PFOA has not been demonstrated to be metabolized, all in vitro assays were conducted both in the presence and in the absence of a mammalian hepatic microsomal activation system. These assays were originally described in twelve contract laboratory reports which have been available via the United States Environmental Protection Agency public docket (Administrative Record 226) for over a decade; however, the details of these assays have not been published previously in the open scientific literature. With the exception of limited positive findings at high and cytotoxic concentrations in some assay trials which reflected the likely consequence of cytotoxic disruption of normal cellular processes and not a specific genotoxic effect, the results of the studies presented in this paper and other published results clearly demonstrate the absence of direct mutagenic or genotoxic risk associated with PFOA. This finding is consistent with the physical/chemical characteristics of PFOA and is supported by other published genotoxicity studies.

a b s t r a c t
Perfluorooctanoate (PFOA) is a fully fluorinated eight-carbon fatty acid analog with exceptional stability toward degradation that has been used as an industrial surfactant and has been detected in environmental and biological matrices. Exposures to PFOA in the workplace and in the environment have continuously stimulated investigations into its potential human health hazards. In this article, the results of fifteen unpublished genotoxicity assays conducted with perfluorooctanoate (as either the linear or linear/branched ammonium salt (APFO) or the linear/branched sodium salt) are reported and include: seven mutation assays (three in vitro reverse mutation assays with histidine auxotrophic strains of Salmonella typhimurium, two in vitro reverse mutation assays with the tryptophan auxotrophic Escherichia coli WP2uvr strain, one in vitro mitotic recombination (gene conversion) assay with Saccharomyces cerevisiae D4, and an in vitro Chinese hamster ovary (CHO) HGPRT forward mutation assay); seven studies to assess potential for chromosomal damage (three in vitro CHO chromosomal aberration studies, an in vitro human whole blood lymphocyte chromosomal aberration study, and three in vivo mouse micronucleus assays); and an in vitro C3H 10T1/2 cell transformation assay. Although PFOA has not been demonstrated to be metabolized, all in vitro assays were conducted both in the presence and in the absence of a mammalian hepatic microsomal activation system. These assays were originally described in twelve contract laboratory reports which have been available via the United States Environmental Protection Agency public docket (Administrative Record 226) for over a decade; however, the details of these assays have not been published previously in the open scientific literature. With the exception of limited positive findings at high and cytotoxic concentrations in some assay trials which reflected the likely consequence of cytotoxic disruption of normal cellular processes and not a specific genotoxic effect, the results of the studies presented in this paper and other published results clearly demonstrate the absence of direct mutagenic or genotoxic risk associated with PFOA. This finding is consistent with the physical/chemical characteristics of PFOA and is supported by other published genotoxicity studies.

Introduction
Perfluorooctanoate (PFOA, CF 3 (CF 2 ) 6 CO 2 − , CASRN 335-67-1) is a perfluorinated carboxylate that has been used principally in the form of the ammonium salt (NH 4 + PFOA or APFO, CF 3 (CF 2 ) 6  emulsifier in the production of fluoropolymers such as polytetrafluoroethylene. The carbon-fluorine bonds impart extreme stability against degradation of the PFOA molecule. The stability of PFOA renders it persistent in environmental and biological matrices. PFOA has been found widely distributed in human blood at low part per billion levels [1,2] and has been identified in wildlife and environmental media [3].
The toxicology of PFOA has been studied extensively and detailed review have been published [4][5][6]. In laboratory toxicology study with mice, exposures to PFOA leads to decrease in body weight, triglyceride, and cholesterol. The serum elimination half-life is approximately 2-3 weeks in either male or female mice and the acute oral LD50 is approximately 450 mg/kg. PFOA is tightly bound to serum proteins and accumulates in the liver, which is the primary target organ for PFOA toxicity. In rodents such as mice, the hepatic changes are susceptible, largely due to nuclear xenosensor receptors PPAR␣ activation which subsequently leads to hepatocellular hypertrophy and proliferation. PFOA can cause developmental delays and neonatal mortalities in mice when pregnant dams are exposed to PFOA during gestation; however, most developmental toxicities appeared to be PPAR␣-mediated [7] and were not observed when PPAR knockout mice were evaluated.
Even though the biological properties of PFOA, including genotoxicity, have been well characterized and reviewed [4][5][6], the details of many key studies conducted in the past that have examined PFOA for potential genotoxicity have not been published. Although the reports for these studies have been made public and available through the USEPA Administrative Record (Docket AR226), for greater accessibility, the experimental details and results of these key studies are summarized and presented herein. Information from other published studies is also included in the discussion to provide a more complete evaluation of perfluorooctanoate for potential genotoxicity.

Materials
Five perfluorooctanoate (PFOA) test materials were used in conducting the genotoxicity studies reported herein. The characteristics of these five samples and the specific studies in which they were used are summarized in Table 1. The purity of the solids used in all samples was ≥95%. Four of the PFOA samples were tested as the ammonium salt (APFO), and one was tested as the sodium salt (Na + PFOA). Three of the APFO samples and the Na + PFOA sample were produced by electrochemical fluorination (ECF) and contained approximately 22% branched isomers of PFOA; for example, as representative lot may contain 12.6% internal monomethyl (non-alpha); 9% isopropyl; 0.2% tert-butyl; 0.1% gem-dimethyl; and 0.1% alpha monomethyl. Impurities in ECF-produced PFOA primarily consisted of lesser homologs (C4-C7) at approximately 2% with additional minor impurities (monohydro APFO, monounsaturated APFO, and undefined (possibly) substituted perfluorocyclo species). One of the APFO samples  [8]. The test methods employed were originally described by Ames et al. [9] and Maron and Ames [10]. Griffith and Long [11] previously provided a brief description of a study in which S. typhimurium histidine auxotrophic strains TA98, TA100, TA1535, TA1537, and TA1538 were used to test APFO for potential mutagenicity (Sample A, Study 1). A more detailed description of methods is provided here and study data not shown by Griffith and Long are presented in the results. Liver S9 extract was isolated from an adult male Sprague Dawley rat after Aroclor TM 1254 treatment. S9 liver extract (0.10 mL) was mixed with triphosphopyridine nucleotide (TPN) sodium salt (4 mol), glucose-6-phosphate (5 mol), sodium phosphate dibasic (100 mol), MgCl 2 (8 mol), and KCl (33 mol) prior to each assay. The study was performed with the test strains both in the presence and absence of S9 activation. Approximately 10 8 cells from an overnight culture of each indicator strain were added to separate test tubes containing 2.0 mL molten agar supplemented with biotin and histidine. APFO concentrations tested ranged from 0.1 g to 500 g. When microsomal activation was required, 0.5 mL of S9 mix was added. The solution was gently mixed and overlaid onto minimal agar plates. After overlay had solidified, the plates were incubated for 48 h at 37 • C for S. typhimurium strains. Vehicle control (50 L DMSO per plate) was used as the negative control for all the assays while the positive controls used for each strain, with or without S9 activation, are listed in Table 2. Plates were evaluated both macroscopically and microscopically, and the number of revertant colonies was counted and reported. Criteria for a positive response were, in general, based on data from at least three non-cytotoxic concentrations, a positive dose-response, and at least a 2-3 fold Table   2 Positive   increase in revertants per plate as compared to the vehicle control at the highest dose. In two additional assays using APFO (Sample D, Study 8) or Na + PFOA (Sample E, Study 12), S. typhimurium strains TA98, TA100, TA1535, and TA1537 were used. The APFO sample assay was run in duplicate; whereas, the Na + PFOA sample was tested once. Following dose range-finding studies, doses of 30-1500 g per plate APFO (Sample D, Study 8) and 20-1000 g per plate of Na + PFOA (Sample E, Study 12) were tested using triplicate plates at each concentration. Deionized water was used as the negative control, and the positive controls used for each strain were as listed in Table 2. When the S9 metabolic activation system was not incorporated, aliquots of each testing strain (100 L) and test material (50 L) were added to 2.5 mL of molten selective top agar maintained at 45 ± 2 • C. When S9 activation was required, 0.1 mL Aroclor TM 1254-treated male Sprague Dawley rat liver S9 extract homogenates (Molecular Toxicology, Inc., Boone, NC) was mixed with water (0.70 mL), 1 M NaH 2 PO 4 /Na 2 HPO 4 , pH 7.4 (0.10 mL), 0.25 M glucose-6phosphate (0.02 mL), 0.10 M NADP (0.04 mL), and 0.825 M KCl/0.2 M MgCl 2 (0.04 mL). Aliquots of each testing strain (100 L), test material (50 L), and S9 mix (500 L) were added to 2.0 mL of molten selective top agar maintained at 45 ± 2 • C. The mixtures prepared above were vortexed and overlaid onto 15 mm × 100 mm agar plates. After overlay had solidified, the plates were inverted and incubated for 52 ± 4 h at 37 ± 2 • C. The mean and standard deviation of the number of revertant colonies per plate were calculated from the individual revertant colony counts from triplicate cultures at each concentration by strain. Criteria for a positive response were as described above.
2.2.1.2. E. coli reverse mutation assay (Sample D, Study 8; Sample E, Study 12). The potential mutagenicity of APFO (Study 8) and Na + PFOA (Study 12) was studied in the E. coli test system using the tryptophan auxotrophic strain WP2uvrA. Methods employed were originally described by Green and Muriel [12]. Preparation of cultures, dose ranges, and enumeration of revertant colonies were as described above for the S. typhimurium assays conducted as part of Studies 8 and 12 (Section 2.2.1.1). The positive control for the metabolic activation system was 2-aminoanthracene (25 g/plate). When the metabolic activation system was not used, 4-nitroquinoline-N-oxide (1 g/plate) was used as the positive control. The mean number of revertant colonies per plate by concentration and strain were counted and recorded as described above, and the criteria for a positive response were also as described above for the S. typhimurium test strains (Section 2.2.1.1).

2.2.2.
Eukaryotic assay systems 2.2.2.1. S. cerevisiae mitotic recombination (gene conversion) assay (Sample A, Study 1). Also described briefly by Griffith and Long [11] was an assay in which APFO (Sample A) was tested for potential mutagenicity via non-specific DNA damage leading to mitotic recombination expressed as gene conversion to produce prototrophic revertants in the auxotophic heteroallelic D4 strain of S. cerevisiae. The assays followed the S. typhimurium procedures described above for Study 1 (See 2.2.1.1). Plates were evaluated both macroscopically and microscopically and the number of convertant colonies per plate were counted and reported. Study data not shown by Griffith and Long are presented in the results.
Plates were seeded with cells and incubated at 37 • C for 24 h followed by exposure to positive controls (EMS for the non-activated and DMBA for the S9-activated), negative control (Ham's F-12 medium), or APFO (9.75-39 g/mL) for 17 h (non-activated) or 5 h (S9 activated). After exposure, cells were washed twice with phosphate buffered saline (PBS) and incubated for additional 48 h (non-activated) or 64.5 h (S9 activated) in Ham's F-12 medium. The cells were subsequently harvested, re-plated, and passed every 48-72 h for 6 days for phenotypic expression. Following this, cells were grown in selective medium specific for mutant cells for 9 days allowing for colony formation. The cells were rinsed with PBS, fixed in methanol, and stained with Giemsa. The results of the assay were evaluated based on the number of thioguanine-resistant mutants per 10 6 surviving cells. The significance of the test results were determined by using the statistical program, Tallarida, R.S. and R.B. Murray's Pharmacological Calculations Procedure, ANOVA, and Newman-Keuls test for confirmation. The statistical method determined if there was a significant (p ≤ 0.05) increase in the mutation frequency of the test article compared to the negative control article. Three test materials were evaluated in three separate studies using CHO cells both in the presence and absence of a metabolic activation system and with varying test material exposure and harvest times, as summarized in Table 3. These were two APFO samples (Sample C, Study 4; Sample D, Study 7) and one Na + PFOA sample (Sample E, Study 11). Positive controls used were mitomycin C (MMC) for assays without metabolic activation and cyclophosphamide (CP) for assays with metabolic activation. Negative controls for cultures contained only cells and medium while the solvent controls employed the highest concentration of solvent used in the test system. All assays were conducted using duplicate cultures, and data based on the average of duplicate cultures was reported. Multiple experimental trials in the absence and presence of a metabolic activation system were performed for Samples D and E (Studies 7 and 11, respectively), while only one trial for each metabolic activation condition was performed for Sample C (Study 4). Assays conducted in the absence of a metabolic activation system were performed as follows. In two studies with APFO, test concentrations of APFO ranged up to 600 g/mL and in the study with Na + PFOA, test concentrations ranged up to 996 g/mL. The MMC positive control concentrations ranged from 0.08 to 1 g/mL. In all cases, cells were cultured for 24 h prior to treatment by seeding approximately 0.8-1.2 × 10 6 cells per 75 cm 2 flask into 10 mL of McCoy's 5a culture media followed by incubation with the test articles at 37 • C for 3-41.8 h (Table 3). Following incubation with test articles, cultures were washed with PBS and incubated with fresh medium containing Colcemid TM (0.1 g/mL) for 2 h, after which the cultures were harvested from 20 to 44.2 h, based on study (Table 3), and fixed with absolute methanol:glacial acetic acid (3:1) fixative, dried, and stained with 5% Giemsa solution for the analysis of the mitotic index and chromosomal aberrations.
Assays conducted in the presence of a metabolic activation system were performed as follows. In the two studies conducted with APFO, tested ranged up to 1500 g/mL (Sample C, Study 4; and Sample D, Study 7), and, for Na + PFOA, concentrations tested ranged up to 1000 g/mL (Sample E, Study 11). Concentrations used for the positive control agent, CP ranged from 5 to 10 g/mL. Cells were seeded at approximately 0.8-1.2 × 10 6 cells per 75 cm 2 flask in 10 mL of McCoy's 5a culture media and were cultured for 24 h prior to treatment, after which cells were incubated for 2-3 h at 37 • C in McCoy's 5a medium without FBS with added test articles and S9 reaction mixture. Treated cells then were washed with PBS and fresh medium and incubated with Colcemid TM (0.1 g/mL) during the last 2 h of incubation. The metaphase cells were harvested after 20-44.2 h, fixed with absolute methanol:glacial acetic acid (3:1), washed, dried, and stained with 5% Giemsa solution for the analysis of the mitotic index and chromosomal aberration.
Stained cells were selected for good morphology, and only cells with the number of centromeres equal to the modal number of 21 ± 1 were analyzed for mitotic index and chromosomal aberrations. A total of 100 cells per dose level from each treatment were analyzed for different types of chromosomal aberrations. The number of mitotic cells per 1000 was reported as the mitotic index, and, in evaluation of chromosomal aberration data, the percent of cells with aberration, the percent of cells with more than 1 aberration, and evidence for increasing amount of damage with increasing dose was considered, along with evidence of polyploidy and endoreduplication.
In these assays, statistical analysis employed the Fisher's Exact Test [13] to compare the percentage of cells with aberrations in each treatment group with the results from the solvent controls. A linear trend test of increasing number of cells with aberrations with increasing dose [14] was also performed. Significance was established when p < 0.01.

Chromosomal aberrations in human whole blood lymphocytes (Sample E, Study 10)
Human venous whole blood from a healthy adult male donor was drawn into a heparinized Vacutainer ® tube. Whole blood lymphocyte cultures were obtained after heparinized blood (0.6 mL) was incubated with RPMI 1640 culture medium (9.4 mL, supplemented with 15% FBS, 100 U/mL penicillin, 100 g/mL streptomycin, 2 mM l-glutamine, and 1% phytohemagglutinin) at 37 • C, and cultures were incubated for 2 days before treatment with test compound. Na + PFOA (prepared in water) was evaluated in the whole blood lymphocyte culture at concentrations ranging from 12.4 g/mL to 600 g/mL. Dose levels of Na + PFOA were selected for the study based on the preliminary range finder data which evaluated cytotoxicity at doses ranging from 0.033 to 1000 g/mL. Fresh RPMI 1640 culture medium was used as the negative control for the assay. CP (20-50 g/mL in water) and MMC (0.1-0.3 g/mL in water) were used as the positive controls for the assay in the presence and absence of S9 metabolic activation, respectively.
In performing the assay in the absence of a metabolic activation system, whole blood lymphocyte culture was initiated for 2 days followed by exposure to Na + PFOA for up to 46 h. The cultures were then washed with PBS followed by incubation with RPMI 1640 medium containing 0.1 g/mL Colcemid TM . Two hours later, the cells were harvested, dried, and stained in 5% Giemsa solution for analysis of chromosome aberrations.
When metabolic activation was used in the assay system, a preparation of rat S9 liver extract (15 L, Molecular Toxicology, Inc., Boone, NC) was mixed with NADP (1.5 mg/mL) and isocitric acid (2.7 mg/mL). Following 3 h of incubation with Na + PFOA or control substances in the presence of the S9 activation system, cultures were washed with PBS, replenished with fresh RPMI 1640 culture medium, and incubated for up to 46 h. The cells were incubated for the rest of the culture period up to the time of harvest with 0.1 g/mL Colcemid TM present during the last 2 h of incubation. The metaphase cells were harvested and the resulting cell pellets were re-suspended with 75 mM KCl hypotonic solution, fixed with absolute methanol:glacial acetic acid (3:1, v/v), and a portion spread onto a microscope slide. After air-drying, the slide was stained with 5% Giemsa solution and evaluated for mitotic index and chromosomal aberrations.
Assessments of chromosomal aberrations and mitotic indices were performed by first selecting cells for appropriate morphology. Only cells with 46 centromeres (normal number) were analyzed. A total of 100 cells per dose level from each treatment were analyzed for different chromosomal aberrations. Evaluation of cultures was performed as noted above the assays with CHO cells (Section 2.3.1).
Cochran-Armitage test for linear trend and Fisher's Exact Test [15] were employed to compare the percentage of cells with aberrations, polyploidy, and endoreduplication. Test article significance was established where p < 0.01.

In vivo mouse micronucleus assays (Sample C, Study 3; Sample D, Study 6; Sample E, Study 9)
Three separate in vivo studies were conducted to assess the potential for PFOA to increase micronuclei in polychromatic erythrocytes isolated from mouse bone marrow. The methods employed were first described by Heddle et al. [16]. For all three studies, adult male and female Crl:CD-1 ® (ICR) BR mice (6-8 weeks old) were purchased from Charles River Laboratory (Portage, MI). All mice were housed in standard cages; rodent chow (Purina ® Certified Laboratory Pellets ® #5002) and tap water were provided ad libitum. Environmental controls for the animal room were set to maintain a temperature of 72 ± 6 • F, humidity of 55 ± 15%, and a 12-h light/dark cycle. The study facility was accredited by the Association for Assessment and for the Accreditation of Laboratory Animal Care International. All procedures involving mice were reviewed and approved by the Institutional Animal Care and Use Committee. Animal care and procedures followed guidelines as specified the U.S. Department of Health and Human Services Guide for the Care and the Use of Laboratory Animals [17].
The three test articles and dosages utilized were as follows: 1. Na + PFOA (Sample E, Study 9) dose levels of 250 mg/kg, 500 mg/kg, and 1000 mg/kg were selected based on the preliminary range finder data which evaluated the acute oral toxicity of Na + PFOA which ranged from 100 to 1000 mg/kg. Na + PFOA solutions (prepared in deionized water) were administered orally as a single dose to groups of mice (n = 5/sex/dose group/time point). CP (80 mg/kg) and deionized water were used as positive and negative controls, respectively. 2. APFO (Sample C, Study 3) dose levels of 200 mg/kg, 400 mg/kg, and 800 mg/kg were selected for the study based on the preliminary range finder data which evaluated the acute oral toxicity of APFO which ranged from 200 to 1000 mg/kg. APFO solutions (prepared in deionized water) were administered orally as a single dose to primary groups of mice (n = 5/sex/dose group/time point). A secondary group of mice (n = 5/sex) were also treated with 800 mg/kg APFO as possible replacements should mortality occur within the primary group. CP (80 mg/kg) and deionized water were used as positive and negative controls, respectively. 3. APFO (Sample D, Study 6) was tested in the same manner as for Sample C (Study 3) except that dose levels of 150 mg/kg, 300 mg/kg, and 600 mg/kg were used (based on the preliminary range finder data which evaluated the acute oral toxicity of APFO ranging from 300 to 900 mg/kg), and the secondary group was dosed with 600 mg/kg APFO.
At designated times (24,48, and 72 h post dose), mice treated with APFO or Na + PFOA were euthanized by CO 2 asphyxiation, and bone marrow was flushed from the bone into a centrifuge tube containing 3-5 mL of bovine serum. Following centrifugation and the removal of supernatant, a portion of the resulting bone marrow pellet was spread onto a microscope slide, dried, and fixed in methanol and stained with May-Grunwald and Giemsa. The mice treated with both the positive and negative control solutions were euthanized at 24 h post dose only and bone marrow samples were also harvested and evaluated as described above. Bone marrow cells were analyzed for the presence of micronuclei in both the polychromatic erythrocyte (PCE), and the mature normochromatic erythocyte (NCE). Approximately 1000 PCEs per mouse were scored and the frequency of micronucleated cells was expressed as % micronucleated cells based on the number of PCEs analyzed. The spontaneous (background) micronuclei frequency was determined based on the number of PCEs observed in the optic fields while scoring the first 1000 erythrocytes on the slide. The normal frequency of micronuclei in this Crl:CD-1 (ICR) BR strain is about 0-0.4%. The ratios of PCE to NCE were calculated.
The analysis of data was performed using ANOVA [18] on either untransformed (when variances are homogeneous) or rank transformed (when variances are heterogeneous) proportions of cells with micronuclei per animal. If ANOVA was significant (p < 0.05), a Dunnett's ttest [19,20] was used to determine which dose groups, if any, were significantly different from the negative control.

Cell transformation assay (Sample B, Study 2)
The potential of PFOA to transform mouse embryoderived C3H 10T1/2 cells was studied in vitro based on methods originally described by Reznikoff et al. [21,22]. Prior to performance of the transformation assay, cytotoxicity data were obtained and expressed as plating (cloning) efficiency. APFO (0.1-20 g/mL) was added to the cultures that were seeded the day before at 300 cells per 60 mm diameter culture dish. After 24-h exposure (6 replicates per dose) in 5 mL Eagle's basal medium supplemented with 10% FBS, APFO was removed with a complete change of media. Media was replenished every 3 days. After 7 days in culture, the plates were washed in PBS, fixed with absolute methanol, and stained with Giemsa. Solvent control (10 g/mL DMSO) and serum baseline controls were included. The number of colonies per plate was counted and the plating efficiency (PE) was determined by the formula: PE = average number of colonies per plate number of cells seeded per plate × 100 The study was conducted in two phases in that colony transformation was evaluated in the first phase while foci transformation was evaluated in the second phase. For the colony transformation assay, six replicates plates seeded with 300 cells per plate were treated with APFO (0.1-200 g/mL), a positive control (benzo(a)pyrene, 0.1-10 g/mL), and a solvent control (10 g/mL DMSO) for 24 h. After removal of the chemicals, culture media was replenished every 3 days. After 14 days, the plates were washed in PBS, fixed with absolute methanol, and stained with Giemsa. Each colony was examined macroscopically and microscopically and scored for transformation according to criteria of Reznikoff et al. [21].
For the foci transformation assay, six replicates plates seeded with 300 cells per plate were treated with APFO (0.1-100 g/mL), a positive control (butadiene diepoxide, 0.0001-0.01 g/mL), and a solvent control (10 g/mL DMSO). Following exposure for up to 38 days, the plates were washed in PBS to remove the chemical, fixed with absolute methanol, and stained with Giemsa. Focal area was examined macroscopically and microscopically and scored for transformation according to criteria of Reznikoff et al. [21].

Results
3.1. Assays to assess potential to cause point mutations 3.1.1. Prokaryotic assay systems 3.1.1.1. S. typhimurium reverse mutation assays (Sample A, Study 1; Sample D, Study 8; Sample E, Study 12). For these three separate assays, samples tested included linear APFO (Sample D, tested up to 1500 g/plate), linear/branched APFO (Sample A, tested up to 500 g/plate), and linear/branched Na + PFOA (Sample E, tested up to 100 g/plate). No evidence for a significant increase in the number revertant colonies was obtained in these assays utilizing several sensitive histidine auxotrophic strains of S. typhimurium at test doses of up to 1500 g/plate, both in the presence and in the absence of a metabolic activation system (Table 4). It should be noted that with Sample D in Trial 2 (see Study 8), when tested in the absence of the metabolic activation system, a non-dose-responsive fivefold increase in mean revertant colonies was observed with S. typhimurium TA1537 at 1000 g/plate when compared to Trial 1. This could be due to the fact the concurrent vehicle control value for mean revertant in Trial 2 was 2 ± 2 while the vehicle control value was higher in Trial 1, which was reported at 5 ± 2. To clarify, a third trial with the TA1537 strain in the absence of the metabolic activation system was conducted and found a lack of mutagenic potential. In addition, these assays evaluated samples with linear APFO (Sample D), linear/branched APFO (Sample A), and linear/branched Na + PFOA (Sample E), it is reasonable to conclude that from these data, PFOA was not mutagenic under the test conditions, regardless of salt form or isomeric composition.

E. coli reverse mutation assay (Sample D, Study 8;
Sample E, Study 12). No evidence for a significant increase in the number revertant colonies was obtained in two separate assays utilizing a sensitive tryptophan auxotrophic strains of E. coli at test doses of up to 1500 g/plate, both in the presence and in the absence of a metabolic activation system (Table 4). Similar to above, the samples tested included both linear APFO (Sample D, tested up to 1500 g/plate) and linear/branched Na + PFOA (Sample E, tested up to 100 g/plate). Again, from these data, PFOA was not mutagenic under the test conditions regardless of salt form or isomeric composition.
3.1.2. Eukaryotic assay systems 3.1.2.1. S. cerevisiae D4 mitotic recombination (gene conversion) assay (Sample A, Study 1). In the D4 strain of S. cerevisiae, no increase in convertant colonies were seen with APFO when tested in the absence or presence of a metabolic activation at concentrations up to 500 g/plate (Table 4). Thus, APFO did not induce non-specific DNA damage leading to mitotic recombination measured as gene conversion in this assay.
3.1.2.2. CHO/HGPRT forward mutation assay (Sample C, Study 5). On the basis of a preliminary range finding assay, dose levels of APFO at 9.75 g/mL, 19.5 g/mL, and 39 g/mL were selected for this assay. While exposure to the test article at these concentrations did not affect the growth with the regular culture medium, they did result in cytotoxicity with 50% cell survival in selective medium with 6-thioguanine ( Table 5). None of the three selected dose levels showed a statistically significant increase in the number of mutant frequencies. Positive controls for non-activation (EMS) or activation (DMBA) showed the expected significant increased frequency in the number of mutant colonies proving the sensitivity of the test system. Under these experimental conditions, APFO is considered non-mutagenic in this forward mutation assay in CHO cells.  Table 3 summarizes the results of the three in vitro chromosomal aberration assays conducted in CHO cells: one with linear/branched APFO (Sample C, Study 4); one with linear APFO (Sample D, Study 7); and one with linear/branched Na + PFOA (Sample E, Study 11).

Assays
In cultures with APFO exposure (Samples C and D) that lacked the metabolic activation system, no significant increases in cells with chromosomal aberrations, polyploidy, or endoreduplication were observed at APFO concentrations up to the maximum for which data could be reasonably assessed, 300 g/mL. Cytotoxicity, based on reduced mitotic index, was evident at concentrations ranging from 100 to 300 g/mL, dependent on assay trial.
For Na + PFOA (Sample E), five assays were conducted in the absence of the metabolic activation system for exposure periods of 3-41.8 h, and harvest times of 20-44.1 h. Maximum concentrations for which data could be analyzed varied from 198 g/mL (41.8-h exposure, 44.1 h harvest) to 748 g/mL (3-h exposure, 44 h harvest). No chromosomal aberrations were noted for cultures that could reasonably be assessed in the absence of significant cytotoxicity. In the assay that incorporated a 3-h exposure period and 44 h harvest, one of the two duplicate cultures at the cytotoxic 748 g/mL concentration was observed to have 11% of cells with aberrations, while the parallel duplicate had 4% of cells with aberrations (a value similar to the negative and solvent control values) yielding an average of 7.5% of metaphase cells with aberrations. These cultures also had 24.5% of polyploid metaphase cells. Evidence for cytotoxicity at the 748 g/mL concentration in these cultures included a 50% reduction in monolayer confluency and a 57% reduction in mitotic index. In the assay incorporating a 3 h exposure and 20 h harvest, it should be    Table 2 for the corresponding positive controls for each strain. noted that the combined duplicate cultures exposed to the 748 g/mL concentration had only 38 metaphase cells available for analysis as opposed to the 200 required by protocol. Although seven metaphase cells were observed to have chromosomal aberrations, the 748 g/mL cultures were excluded from analysis due to severe toxicity, and the chromosomal aberrations were attributed to the obvious high toxicity at 748 g/mL. No increase in polyploid cells was evident in the assays conducted in the absence of metabolic activation, with the exception of one of the two duplicate cultures at 748 g/mL. Therefore, Na + PFOA was not considered clastogenic in the absence of severe cytotoxicity. When tested in the presence of the metabolic activation system, no increase in chromosomal aberration was observed after linear/branched APFO exposure (Sample C) at concentrations up to the maximum concentration for which data could be assessed, 750 g/mL. Similarly, 3-h exposure to linear APFO (Sample D) in the presence of the metabolic activation system with a 20 h harvest time did not produce cytogenetic effects in the initial trial at concentrations up to the maximum analyzed, 750 g/mL. However, increases over control in percent cells with aberrations were observed in two confirmatory trials with Sample D (3-h exposures and 20.1 h and 44.2 h harvest times) at the highest concentrations for which data could be assessed in each trial (12.5% and 13% at 750 g/mL and 825 g/mL, respectively, for the 20.1 h and 44.2 h assays, respectively). These findings were accompanied by cytotoxicity as evidenced by an increase in the percent of cells with endoreduplication at 675 g/mL and higher concentrations and effects on monolayer confluence and mitotic index. In the 20.1 h assay, for the 600, 675, and 750 g/mL concentrations, monolayer confluence was reduced by 15, 30, and 45%, respectively, and mitotic index by 15, 11, and 30%, respectively. Similarly, in the 44.2 h assay, for the 675, 750, and 825 g/mL concentrations, monolayer confluence was reduced 15, 45, and 55%, respectively, with mitotic index reductions of 17, 56, and 53%, respectively. Floating dead cells and debris were also noted at 750 and 825 g/mL.
When Na + PFOA was tested in the presence of the metabolic activation system in three assay trials, no increase in chromosomal aberrations or polyploid cells was noted at concentrations up to 500 g/mL; however, contradictory results were observed at 748 g/mL in two replicate assay trials in which cells were exposed 3 h with a 20 h harvest time. At that concentration, a statistically significant percent of cells with chromosomal aberrations (14.5%) and percent polyploid cells (7%) occurred in one of the two replicate assay trials but not in the other. The highest concentration tested in these two replicate assay trials, 1000 g/mL, was clearly cytotoxic, and increased percent cells with aberrations (31.5% and 42.7%) and increased percent polyploid cells (15% and 26%) were statistically significant over control in both replicate assay trials at this concentration. Cytotoxicity at 1000 g/mL was severe, consisting of large reductions in monolayer confluency (70% and 95%) and mitotic index (14% and 47%). In the single assay trial with 3-h exposure period and 44.1 h harvest time, no increase in the percent of cells with either chromosomal aberrations or polyploidy was observed at 748 g/mL, the highest concentration that allowed for analysis. At the highest concentration tested under these exposure and harvest conditions, 1000 g/mL, mitotic index at was reduced by 70%, disallowing analysis.
The observation of an increase in chromosomal aberration and polyploidy with Sample D at cytotoxic concentrations in the presence of a metabolic activation system are of questionable relevance for the assessment of the clastogenicity of APFO. These effects were not noted in the absence of the metabolic activation system or in the absence of cytotoxicity. The physical/chemical properties of APFO preclude metabolic activation, DNA binding, and intercalation. Therefore, APFO was not considered to be a selective genotoxic agent either in the presence or in the absence of a metabolic activation system. Similarly, the observed chromosomal aberrations and polyploidy at the highest concentrations of Na + PFOA (Sample E) was considered to be the likely consequence of severe cytotoxicity leading to disruption of normal cellular processes and not a specific genotoxic effect of PFOA [23,24].  3.2.1.2. Chromosomal aberrations in human whole blood lymphocytes (Sample E, Study 10). Presented in Table 6 are data from the chromosomal aberration assay using human whole blood lymphocytes. The test compound produced cytotoxicity at concentrations above 100 g/mL in the absence of the metabolic activation system and above 200 g/mL with the metabolic activation system. Regardless of S9 activation, Na + PFOA treatments caused a dose-dependent reduction in mitotic index indicative of toxicity. The extent of % reduction was very pronounced at higher Na + PFOA doses. Even so, when chromosomal aberrations were analyzed from cultures treated with higher concentrations of Na + PFOA doses, there were no statistically significant increases in the percent of cells with chromosomal aberrations or polyploidy, when compared to the cells treated with vehicle control. Endoreduplications were generally not observed following treatment with Na + PFOA with the exception of one instance at highly toxic dose level at which the mitotic index was reduced by about 80%. This effect was considered to be the consequence of generalized cell damage as opposed to selective genotoxicity, because it occurred at dose levels for which it was not possible to assess a sufficient number of surviving metaphase cells in one of the duplicate cell cultures. Cells treated with positive controls had statistically significant increases in the percent cells with aberrations in the absence or presence of metabolic activation.

3.2.2.
In vivo mouse micronucleus assay (Sample C, Study 3; Sample D, Study 6; Sample E, Study 9) Presented in Table 7 are data for percent micronucleated PCEs and the ratio between PCE and NCE from three separate studies, one with Na + PFOA (Sample E, Study 9) and two with APFO (Sample C, Study 3 and Sample D, Study 6). No significant increases in bone marrow PCE micronuclei occurred in mice treated with the test compounds in any of the three studies. On one occasion, in females treated with 800 mg/kg APFO at the harvest time 48 h post dosing, the rate of micronucleated PCE was slightly increased over vehicle control (0.3%). This singular observation is considered not biologically relevant because: (1) it was within the historical control range, (2) it was not observed at a later harvest time (72 h post dose) in the same experiment, (3) it was restricted to one sex, and (4) it was not observed at an even higher dose in females treated with 1000 mg/kg of Na + PFOA.
In mice treated with cyclophosphamide, the positive control, both sexes had a statistically significantly increase in the percentage of micronucleated cells. Male and female mice dosed with 1000 mg/kg of Na + PFOA had a statistically significant decrease in PCE/NCE ratio, indicating bone marrow toxicity and demonstrating that the test compound had reached the target organ.

Cell transformation (Sample B, Study 2)
The data for the C3H 10T1/2 cell transformation assay are presented in Table 8. While APFO showed a dose-dependent decrease in colony formation and plating efficiency, there was no morphologic evidence of Table 7 In vivo mouse micronucleus data summary table, data are presented as mean ± standard error. Adult male and female Crl:CD-1 ® (ICR) BR mice were treated with a single oral dose of test compound with n = 5/sex/time point unless specified otherwise (Studies 3, 6, and 9). transformation in the colonies treated with APFO. A longer term foci transformation regimen was also evaluated for APFO and there was no evidence of foci or transformed foci. The response to positive controls (benzo(a)pyrene and butadiene epoxide) confirmed the sensitivity of the assay as expected, and all findings observed in positive controls were absent in solvent controls. In summary, APFO did not induce cell transformation in this in vitro system.

Discussion
The potential genotoxicity of PFOA was tested in fifteen assays covering several genotoxicological endpoints. These assays included: seven mutation assays (three in vitro reverse mutation assays with histidine auxotrophic strains of S. typhimurium, two in vitro reverse mutation assays with the tryptophan auxotrophic E. coli WP2uvr strain, one in vitro mitotic recombination (gene conversion) assay with S. cerevisiae D4, and an in vitro Chinese hamster ovary (CHO)/HGPRT forward mutation assay); seven cytogenetic aberration studies (three in vitro Chinese Hamster Ovary chromosomal aberration studies, an in vitro human whole blood lymphocyte chromosomal aberration study, and three in vivo mouse micronucleus assays); and an in vitro C3H 10T1/2 cell transformation assay. Although these studies have been available as laboratory reports via the United States Environmental Protection Agency Administrative Record 226 (AR-226, see Table 1) and briefly summarized in previous reviews [5,6], the details of these assays have not been published. The results of these assays demonstrate no significant potential for genotoxicity, regardless of PFOA salt form (ammonium or sodium) or isomeric composition (combination of linear and branched or pure linear).
Many of the assays reported incorporated a metabolic activation system. It should be noted that metabolism of PFOA by microsomal systems has never been demonstrated and would not be expected based on the exceptional strength of the carbon-fluorine bonds in the alkyl chain  [25][26][27][28][29]. Therefore, formation of an active metabolite by exposure of PFOA to the microsomal activation system is precluded. However, it is worth noting that the S9 proteins included in the metabolic activation system may sequester PFOA through ionic binding [30]. PFOA has not shown the potential to induce DNA point mutations or mitotic recombinations in a series of assay systems. These included the bacterial reverse mutation assays utilizing S. typhimurium histidine auxotrophic strains (TA98, TA100, TA1535, TA1537, and TA1538) and E. coli tryptophan auxotroph (WP2uvr), the S. cerevisiae D4 mitotic gene conversion/recombination assay, and the Chinese hamster ovary cell (CHO) forward mutation assay. Three independently published studies utilizing various S. typhimurium strains exposed to PFOA in the presence or absence of a metabolic activation system were consistent with the results reported herein. In one published study, PFOA was not mutagenic in four strains of S. typhimurium (TA98, TA100, TA102, TA104) when tested at concentrations up to 500 M (206 g/mL) [31]. In another study by Endo et al. [32], S. typhimurium strains TA 98 and TA100 exposed to PFOA at unspecified concentrations did not show an increase in revertant colonies. Using the assay for umuC gene expression assay with S. typhimurium TA1535/pSK1002 (hisG46, rfa, uvrB) to analyze for ␤galactosidase activity, no significant increased activity was seen at concentrations of PFOA up to 1000 M (413 g/mL), indicating a lack of mutagenicity in the assay system [33].
The studies reported herein also demonstrated that PFOA lacked the potential to produce cytogenetic changes, including chromosomal damage, aberrations and polyploidy under non-cytotoxic exposure conditions. These studies included in vitro assays to assess the potential of PFOA to produce chromosomal aberrations in cultured human lymphocytes and CHO cells, as well as the in vivo assessment of micronucleated polychromatic erythrocytes in PFOA-treated mice. In CHO cells, aberrations were observed only in the presence of significant cytotoxicity in three of 16 assay trials that were conducted. PFOA does not possess the chemical/physical characteristics of a genotoxic agent, is incapable of metabolism [25][26][27][28][29], is not expected, based on its structure, to bind covalently to DNA or to intercalate into the double helix, and lacks the planar characteristics of an intercalating agent. Therefore, it is reasonable to assume that the positive findings at high and cytotoxic concentrations in some CHO chromosomal aberration assay trials were the consequence of cytotoxic disruption of normal cellular processes and not a specific genotoxic effect. This is supported by the general lack of genotoxicity in the in vitro chromosomal aberration study in human lymphocytes, in which no evidence of an increase in chromosomal aberrations, polyploidy, or endoreduplication was observed. Previously published reports in which HepG2 cells were exposed in vitro to PFOA have provided mixed results with respect to the production of DNA strand breaks and micronuclei [34][35][36]. Florentin et al. [35] observed no increases in HepG2 cell DNA strand breaks or micronuclei at PFOA concentrations up to 800 M (330 g/mL), and Eriksen et al. [34] also found no increase in PFOA-induced DNA strand breaks in this cell line at concentrations up to 400 M (165 g/mL). By contrast, Yao and Zhong [36] observed increases in micronuclei in HepG2 cells exposed to 100-400 M (41-165 g/mL) PFOA and increases in DNA strand breaks at concentrations of 50-400 M (21-165 g/mL). Among the assays reported here, the lack of increases in mouse bone marrow micronucleated polychromatic erythrocytes found in the three separate in vivo mouse micronucleus assays with respective oral doses of 600, 800, and 1000 mg/kg provides compelling evidence for a lack of potential for PFOA to induce cytogenetic alteration. Taken together, these assays demonstrate a lack of potential for chromosomal effects as a result of PFOA exposure.
Using the C3H 10T1/2 cell culture system in vitro, APFO did not produce cell transformation at concentrations up to the maximum tested, 200 g/mL. Cell transformation, whether mutational or epigenetic, alters the normal properties, organization, and propagation of cells in culture with emphasis on the acquisition of properties typical of cancer cells. One additional report [37], published previously, has examined to potential of PFOA to produce cell transformation in Syrian hamster embryo (SHE) cells. PFOA tested up to 300 M (124 g/mL) did not induce SHE cell transformation frequency. No genotoxicity in SHE cells was seen via the COMET assay, nor was DNA breakage increased in benzo-␣-pyrene initiated cells. The data reported herein and that reported by Jacquet et al. [38] demonstrate a lack of cell transformation potential in vitro to be associated with PFOA.
The data reported herein are designed to observe for genotoxic effects under non-cytotoxic exposure conditions to PFOA; however, there are several published articles that have reported cytotoxic disruption of cellular homeostasis, modified gene expression, and increased intracellular ROS production as a consequence of in vitro exposure to PFOA. The potential for PFOA to induce oxidative stress, which may relate to potential cytotoxicity and genetic alteration, has been investigated in several in vitro studies and at least one in vivo study. Freire et al. [31] reported an increase in PFOA-induced reactive oxidative species (ROS), as measured by oxidation of 2 ,7 -dichlorofluorescein diacetate (DCFH-DA) to 2 ,7dichlorofluorescein (DCF) in the monkey kidney-derived Vero cell line at the highest concentration tested 500 M (210 g/mL); however, cytotoxicity in these cells was evident at concentrations greater than 50 M (21 g/mL) with dose-related mitochondrial changes as fission and reduced cell density and increased apoptosis. Using HepG2 cells exposed to PFOA, Yao and Zhong [36] reported an observed increase in ROS (measured as DCF fluorescence) as well as 8-hydroxydeoxyguanosine (8-OHdG) DNA adducts. They reported micronuclei breaks occurred with PFOA concentrations at 100-400 M (42-168 g/mL) and DNA strand breaks at 50-400 M (21-168 g/mL). Also using HepG2 cells, Hu and Hu [39] found that PFOA concentrations from 50 to 200 M (20-82 g/mL) induced production of ROS, dissipation of mitochondria membrane potential, and apoptosis. Cellular antioxidant defenses were reduced, including glutathione content, as well as activities of catalase, superoxide dismutase, and glutathione reductase. Differential expression of genes such as p53, Bcl-2, caspase-9 was evident. The authors suggested that these effects occurred via a mechanism in which the homeostasis of antioxidant systems was overwhelmed, thus modifying expression of genes involved with the apoptotic regulation. In another experiment with HepG2 cells, Eriksen et al. [34] found that PFOA increased the intracellular ROS production by 1.52 fold in a non-concentration-dependent manner and did not cause DNA strand breakage and acid-labile sites or an increase in formamidopyrimidine-DNA-glycolase sites; thus, the authors concluded that PFOA, along with other perfluorinated chemicals, induced only modest effects in terms of ROS production and produced no DNA damage in HepG2 cells. Florentin et al. [35] also exposed HepG2 cells to PFOA for 24 h and observed cytotoxicity at concentrations of 200-800 M (83-330 g/mL); however, no increases in ROS, DNA strand breaks, or micronuclei were observed. Following a single intraperitoneal injection of 100 mg/kg PFOA, Takagi et al. [40] found increased liver weights and increased liver 8-OHdG adducts of male Fisher 344 rats. It should be noted that the level of 8-OHdG adducts reported was a marker for oxidative DNA damage only and not a measurement of PFOA-DNA adduct.
Because PFOA is a strong organic acid with excellent surfactant and surface tension-lowering properties, it is likely that some of the biochemical changes observed in vitro, as reported by others above, were mediated by cytotoxicity and surfactant-induced membrane disruption. For perspective on cytotoxicity of PFOA, PFOA has been shown to be similar to naturally occurring fatty acids as well as other perfluorinated carboxylates with respect to having low cytotoxicity in human colon carcinoma (HCT116) cells. For PFOA, the 24-h EC 50 and 72-h EC 50 were 937 M (387 g/mL) and 313 M (130 g/mL), respectively, while the 72-h EC 50 's for the naturally occurring fatty acids decanoic, dodecanoic, and octadecanoic acids ranged from 677 M (279 g/mL) to 2907 M (1200 g/mL) [41]. Therefore, these inconclusive observations are difficult to translate to in vivo conditions and they do not contradict the conclusion that PFOA has not shown the potential for genotoxicity under non-cytotoxic conditions.
When considering the structural and chemical characteristics of PFOA relative to what is known about direct mutagens and clastogens, it is evident that PFOA does not possess structural or chemical properties that would be consistent with those known to be important in identifying potential genotoxic agents. The complete fluorination of the seven carbons of the chain that supports the carboxyl group precludes chemical reactivity under physiological conditions. Neither is PFOA a planar molecule capable of intercalation into DNA. Interactions of PFOA with DNA using circular dichroism indicated a change in DNA conformation with the binding to DNA being non-covalent in which PFOA aligned along the backbones and interacted with the homolateral bases via hydrophobic interactions [42]. DNA binding was not found when tested at either 50 (21 g/mL) or 500 M (210 g/mL) concentrations [43]. In addition, no significant oxidative DNA damage was produced in primary rat testicular cells [44] and Ddit3 gene expression (marker for DNA damage) was unaltered by PFOA in HepG2/C3A human hepatoma cells at concentrations below 200 M (83 g/mL) [45].
In vivo and in vitro mutagenicity/genotoxicity tests are useful predictive tools for evaluating potential genotoxic carcinogens and mutagenicity of chemicals. The results reported in this paper are consistent with the carcinogenicity data obtained from two chronic dietary studies, in which daily dietary treatment with branched/linear APFO up to 300 ppm in the diet of Sprague Dawley rats up to 104 weeks did not produce any excess malignant (invasive) tumors of any type when compared to the control [46,47]. From a mechanistic perspective, the benign tumors observed in these cancer bioassays have been proposed to be most likely the result of non-genotoxic processes related to activation of the xenosensor nuclear receptors in rats such as NR1C1 (PPAR␣) [48][49][50][51].
Therefore, with the exception of limited positive findings at high and cytotoxic concentrations in some CHO chromosomal aberration assay trials which reflected the likely consequence of cytotoxic disruption of normal cellular processes and not a specific genotoxic effect, at non-cytotoxic conditions, the results of the studies presented in this paper and other published results clearly demonstrate the absence of direct mutagenic or genotoxic risk associated with PFOA.

Conflict of interest
John Butenhoff and Shu-Ching Chang are employees of the 3M Company, a former manufacturer of PFOA. Gerald Kennedy is a consultant to DuPont Company, a previous user and manufacturer of PFOA. Reinhard Jung is a consultant to Dyneon, a subsidiary of 3M Company. 3M Company funded the studies reported herein.


extfieldsofstudy: Medicine Biology

s2fieldsofstudy: Biology Medicine

text:
Autophagy is essential for effector CD8 T cell survival and memory formation

The importance of autophagy in memory CD8 T cell differentiation in vivo is not well defined. We show here that autophagy is dynamically regulated in virus-specific CD8 T cells during acute lymphocytic choriomeningitis virus infection. Autophagy decreased in activated proliferating T cells, and was then upregulated at the peak of the effector T cell response. Consistent with this model, deletion of the key autophagy genes Atg7 or Atg5 in virus-specific CD8 T cells had minimal effect on generating effector cells but greatly enhanced their death during the contraction phase resulting in compromised memory formation. These findings provide insight into when autophagy is needed during effector and memory T cell differentiation in vivo and also warrant a re-examination of our current concepts about the relationship between T cell activation and autophagy.

CD8 T cells provide protection against intracellular bacterial, parasitic, and viral infections, as well as cancers 1,2 . Following antigen stimulation, naive CD8 T cells go through many rounds of proliferation, giving rise to effector T cells, which are efficiently eliminate infected cells. Upon antigen clearance, the vast majority of effector CD8 T cells undergo apoptosis, leaving only a small pool of cells to survive and differentiate into memory cells 3,4,5 . During this naive to effector to memory differentiation process, T cells undergo cellular and metabolic reprogramming shifting from anabolic processes and proliferation to catabolic processes and contraction of cell populations to generate memory. It is important to define the role of macroautophagy (herein, "autophagy") during this process.
Autophagy is an evolutionarily conserved process involving the engulfment and delivery of cytosolic contents to the lysosome for degradation 6,7,8,9,10 . This catabolic activity of autophagy is essential for cellular homeostasis and has been suggested to be inversely correlated with cell growth and proliferation 11 . In contrast to this paradigm, it has been reported that autophagy is up-regulated in proliferating T cells 9,12,13 . T cell receptor (TCR) stimulation promotes activation and proliferation of T cells and also induces the metabolic checkpoint kinase mTOR signaling which would be expected to inhibit rather than induce autophagy 8 . Thus, major questions remain related to why and how proliferating T cells upregulate autophagy in the presence of positive mTOR signals when cells need more proteins and organelles to donate to daughter cells. Furthermore, because autophagy has been predominantly studied in vitro during T cell activation little is known about in vivo autophagy activity in antigen specific T cells during the course of effector and memory T cell differentiation after viral infections.
The in vivo functional role of autophagy in antigen specific T cells during viral infections remains unclear, but is important as pharmacologic manipulation of autophagy is being considered for many human diseases 14 . Conditional knockout mice in which either of the key autophagy genes Atg5 or Atg7 genes were selectively deleted during early T cell development using Lck-cre decreases mature peripheral T cells 10,15 . Similarly, reduced peripheral T cells were observed in Atg5 −/− chimeric mice after transplantation of fetal liver cells from Atg5 −/− mice 9 . This study also showed that Atg5-deficient T cells exhibited a reduced proliferative capacity following in vitro TCR stimulation 9 . While these data indicate that autophagy plays a key role in T cell development and homeostasis, they shed less light on the function of autophagy genes in T cells responding to antigen because the cells studied had developed in the absence of autophagy genes such as Atg5 or Atg7 and exhibit abnormalities in gene expression and mitochondrial numbers and function 10,15 . Thus, a new approach using phenotypically normal naive T cells is required to study the functional role of autophagy during T cell activation in vivo.
Here, we examined two important questions; the kinetics of autophagy activity and the role of autophagy during response to lymphocytic choriomeningitis virus (LCMV). In contrast to previous observations, we found that autophagy was inversely correlated to T cell proliferation and was significantly inhibited during the early effector stage (i.e., when the cells are highly proliferative). Autophagy activity was then up-regulated at the peak of expansion (i.e., when antigen is cleared and antigen-specific cells about to undergo contraction phase). We also established the significance of autophagy proteins for the development of CD8 T cell memory in vivo by knocking out either one of the two essential autophagy genes, Atg5 and Atg7 using granzyme B cre system in which normal naive T cells were developed and autophagy genes were deleted only after T cells were activated with antigen. Our study provides important insight into the kinetics and functional role of autophagy in antigen specific CD8 T cells during effector and memory differentiation.

Dynamic regulation of autophagy in virus specific T cells
During an acute viral infection, naive CD8 T cells undergo vigorous clonal expansion, followed by contraction, in which a small percentage of effector cells survive to establish memory (Fig. 1a). To study autophagy in antigen-specific CD8 T cells through the distinct phases of the T cell response, we took several different approaches to analyze the autophagy pathway and autophagic flux in antigen-specific CD8 T cells following acute infection with lymphocytic choriomeningitis virus (LCMV) Armstrong strain. We isolated transgenic CD8 T cells that recognize the LCMV GP33-41 peptide (P14 cells) at distinct stages of the T cell response: the expansion phase; i.e., cells with a blasting phenotype (day 5 post infection (p.i.) when virus specific effector CD8 T cells are actively proliferating); the peak of expansion (day 8 p.i. when effector CD8 T cells stop proliferating and decrease in cell size); the contraction phase (day 15 p.i.); and the memory phase (day 30 p.i.) ( Fig. 1a and b). To assess autophagy activity, we first examined the amount of microtubule-associated protein light chain 3 beta-I (LC3b-I) and LC3b-II in the sorted P14 cells. LC3b-II, the lipidated form of LC3b-I, is a classic marker for autophagosomes, and is incorporated into the elongating membrane that eventually forms the autophagosome and is subsequently degraded after delivery to the lysosome 7 ( Supplementary Fig. 1a). The quantities of both LC3b-I and LC3b-II in antigen-specific CD8 T cells peaked at day 5 p.i. (Fig. 1c). The accumulation of LC3b proteins in P14 cells without an increase in the level of LC3b transcripts at day 5 p.i. (Supplementary Fig. 1b) would be consistent with either an induction of autophagy and increased production of these proteins or an inhibition of lysosomal degradation of these proteins.
To further assess autophagy activity, we therefore examined p62 (also known as Sequestosome-1) protein expression. p62 is a widely used marker for measuring autophagic activity since it is both an autophagy substrate and acts as an adaptor in targeting ubiquitinated proteins for lysosomal degradation via the autophagy pathway 18 ( Supplementary Fig. 1a). Similar to LC3b, p62 protein showed the highest accumulation at day 5 p.i., which was not associated with an increase in the corresponding mRNA ( Fig. 1c and Supplementary Fig. 1c). In addition, we examined if LC3b and p62 proteins were accumulated at a much earlier stage of T cell activation. In these experiments, LCMV GP33-41 peptide was injected into P14 transgenic mice to activate all P14 cells in a short synchronized timeframe. Similar to what was observed in the activated P14 cells obtained from day 5 p.i., we detected higher amounts of both LC3b and p62 proteins at 20 hours post activation without an increase of their mRNAs (Supplementary Fig. 1d and e). Together these data suggested an altered flux in the autophagy pathway at the early effector stage of the T cell response, and were most consistent with a reduction of autophagy during antigendriven T cell activation in vivo.
To further define how the highly dynamic autophagy pathway is regulated in vivo during the CD8 T cell response, we employed a reporter system for autophagy activity by fusing a recombinant mouse LC3b protein to green fluorescent protein (GFP) (Fig. 2a) 7 . This fusion gene was introduced into P14 cells with retroviral vector system, MSCV-IRES-Thy1.1 (MIT), by which MIT-transduced P14 cells can be distinguished by the expression of Thy1.1 ( Fig. 2a and Supplementary Fig. 2a). This reporter system allowed us to quantify autophagy activity by simply measuring GFP expression by flow cytometry; high autophagy activity was reflected by the reduction in GFP intensity due to either the enhanced degradation of GFP-LC3b protein through the autophagy pathway or fluorescence quenching of GFP by low pH after fusion of autophagosomes with lysosomes 19 . As a control we used a G120A mutant of GFP-LC3b, since this mutation removes the C-terminal amino acid to which phosphatidylethanoloamine is conjugated during the generation of LC3b-II from LC3b-I, resulting in this molecule failing to be incorporated into autophagosomes (Fig. 2a). Using this reporter system, we assessed autophagy activity at different times following LCMV infection. Consistent with our analysis of LC3b and p62, at day 5 p.i., GFP from either the WT or mutant LC3b was comparable, indicating lower autophagic flux (Fig. 2b). Similar amounts of GFP expression between WT and mutant LC3b were also observed at much earlier times post infection (days 2 and 3) ( Supplementary Fig. 2b-e). However, at day 8 p.i., in contrast to days 2 to 5, there was a profound difference between the WT and mutant LC3b proteins; we observed a drastic increase in the percentage of GFP-negative P14 cells in the WT relative to the G120A mutant, suggesting a significantly elevated flux through the autophagy pathway at day 8 compared to day 5 ( Fig. 2b and c). This profound difference in GFP intensity between the WT and mutant LC3b groups remained through to day 30 p.i. (Fig. 2b and c). In addition to this P14 transgenic system, we observed a similar change in the autophagy flux in endogenous LCMV specific tetramer-positive CD8 T cells (Supplemental Fig. 3). To apply the reporter system to endogenous cells, hematopoietic stem cells (HSCs) were transduced with MIT containing either WT or G120A mutant GFP-LC3b, followed by adoptive transfer of these HSCs into lethally irradiated mice ( Supplementary  Fig. 3a). After reconstitution, mice were infected with LCMV Armstrong and then GFP expression was monitored in MIT-transduced D b GP33 tetramer-positive CD8 T cells ( Supplementary Fig. 3a). Similar to the results obtained from P14 transgenic T cells, GFP expression in D b GP33 + cells transduced with MIT carrying WT LC3b were comparable to those with MIT carrying G120A mutant at day 6 p.i., and a significant decrease in WT GFP-LC3b expression was observed at days 8, 15 and 30 p.i. (Supplementary Fig. 3b). Taken together, these data indicate that autophagy is decreased in antigen-specific CD8 T cells during the clonal expansion phase, but is significantly increased following the expansion phase and into the memory phase.
As an additional assessment of autophagy, we visualized GFP-LC3b puncta using multispectral imaging flow cytometry (ImageStream ® ) to quantify the number of LC3-positive autophagosomes; GFP puncta reflected the co-localization of lipidated GFP-LC3b incorporated onto the elongating phagophore membrane that subsequently forms the autophagosomes. On days 3, 5, and 8 p.i., we observed puncta in P14 cells transduced with WT GFP-LC3b, although a slightly higher frequency of puncta was detected on day 3 p.i. (Fig. 2d and e). These data indicate that antigen specific CD8 T cells can form autophagosomes during the T cell expansion phase. Collectively our results suggest that the low autophagy activity during clonal expansion is due to reduced flux through the lysosomal degradation pathway and that upon conclusion of the clonal expansion phase (day 8 p.i.) autophagic flux is significantly increased.
Our results concerning the in vivo kinetics of autophagy activity in virus specific CD8 T cells were unexpected because prior in vitro studies showed that T cells increased autophagy after T cell receptor (TCR) stimulation 6,7,9,12,13 . To further confirm our findings and examine if autophagolysosome formation is indeed inhibited during the clonal expansion phase, we made retrovirus vectors by fusing mCherry fluorescent protein into the GFP-LC3b protein used above (Fig. 3a). Since mCherry shows more stable fluorescence compared to GFP under acidic condition such as following fusion of autophagosomes to lysosomes 7, 20, 21 , comparing the amount of GFP fluorescence relative to mCherry allowed us to measure whether LC3b had been delivered from autophagosomes into acidic autophagolysosomes. In this system, enhanced autophagic flux results in reduced GFP fluorescence compared to mCherry fluorescence due to delivery of the LC3b protein into an acidic environment 7, 20, 21 . Using this system, we examined autophagolysosome formation in antigen specific CD8 T cells during the T cell expansion phase (day 5) and at the peak of T cell responses (day 8) (Fig. 3b). Consistent with our previous results, proliferating day 5 effector CD8 T cells were larger in size compared to effector CD8 T cells at day 8 p.i. when these cells had stopped proliferating (Fig. 3c). The GFP/mCherry fluorescence intensity ratio in retrovirus-transduced P14 cells with WT LC3b was very similar to that in P14 cells with G120A mutant LC3b at day 5 p.i. (Fig. 3d and e), indicating that autophagy decreased in proliferating virus specific CD8 T cells. In contrast, at day 8 post infection, GFP fluorescence was specifically decreased compared to mCherry fluorescence in the WT LC3b group (Fig. 3d) and the GFP/mCherry fluorescence intensity ratio showed a 4.3-fold decrease between WT and G120A groups (Fig. 3e). These results are consistent with increased delivery of LC3b into autophagolysosomes due to induction of autophagy between 5 and 8 days after infection. Taken together (Figs. 1-3, Supplementary Figs. 1-3), autophagic flux is inhibited during T cell clonal expansion phase via impairing autophagosome maturation into autophagolysosomes, followed by a rapid induction of autophagy concurrent with the peak of CD8 T cell effector responses that occurs at day 8 post infection.

Atg7 and Atg5 are critical for CD8 T cell memory formation
To determine the importance of the increased autophagy activity during the transition phase from effector to memory CD8 T cells, we generated conditional knockout mice by crossing Atg7 fl/fl or Atg5 fl/fl mice to transgenic mice carrying the Cre recombinase gene driven by the granzyme B promoter (Gzmb-Cre) 22,23,24 . In this system, the critical autophagy genes, Atg5 or Atg7, are specifically deleted upon expression of the granzyme B gene in activated CD8 T cells. Herein, we refer to these genes as Atg7 fl/fl Gzmb-Cre and Atg5 fl/fl Gzmb-Cre.
We infected Atg7 fl/fl Gzmb-Cre mice with the LCMV Armstrong strain. In order to establish the efficiency of the Gzmb-Cre system in the context of Atg7 fl/fl Gzmb-Cre mice, we isolated D b GP33 + CD8 T cells 8 days following infection ( Supplementary Fig. 4a). Assessment of gene deletion using quantitative PCR showed a greater than 20-fold reduction in the genomic copies of the Atg7-flox allele in the D b GP33-specific CD8 T cells, demonstrating a 95% deletion of the floxed allele (Fig. 4a). A similar level of Atg7-flox gene deletion was also observed in the bulk activated CD8 T cell (CD44 hi CD62L lo ) population, >95% of which were LCMV-specific and expressing granzyme B at day 8 p.i. 25 (Fig. 4a and Supplementary Fig. 4b). The extent of Atg7 gene deletion was also reflected by the Atg7 protein within the bulk activated CD8 T cells, whereby Atg7 protein was hardly discernible in the CD44 hi CD62L lo effector CD8 T cell population from Atg7 fl/fl Gzmb-Cre mice. We confirmed the deletion of Atg7 protein by examining its catalytic activity. Atg7 catalyzes reactions resulting in Atg5-Atg12 conjugation and lipidation of LC3-I to give rise to LC3b-II 6 . The enzymatic activity of Atg7 in the effector CD8 T cells was significantly diminished, as indicated by the absence of both Atg5-Atg12 conjugation, as well as lipidated LC3-II ( Fig. 4b). Instead, we detected Atg5 protein in abundance as an un-conjugated species (Fig.  4b). Furthermore, p62 accumulated in the absence of Atg7, indicative of defects in autophagic activity (Fig. 4b). Together, these data indicated the high efficiency of the Cre-loxP system in deleting the target Atg7-flox allele in LCMV-specific CD8 T cells and the consequent loss of autophagy at a time point (day 8) when autophagy would normally be increased. In addition, these data validate the assays of LC3b and p62 used above to measure autophagic flux in primary T cells in vivo.
To assess the importance of Atg7 in antigen-specific CD8 T cells, we infected Atg7 fl/fl Gzmb-Cre mice with the LCMV Armstrong strain and followed the differentiation of LCMVspecific CD8 T cells over time. Eight days after infection, D b GP33 + CD8 T cells had undergone significant expansion in both Atg7 fl/fl and Atg7 fl/fl Gzmb-Cre mice (Fig. 4c). At day 8 p.i. the memory precursor population, characterized by KLRG1 lo and CD127 hi , was present in Atg7-deficient D b GP33-specific cells at similar proportions compared to wildtype cells (Fig. 4c) 3,4 . Other phenotypic markers, including CD44 and CD62L, showed no considerable differences between the two groups ( Supplementary Fig. 4c). Despite a small reduction in PBMC at day 8 p.i. (Fig. 4c left panel), antigen-specific CD8 T cells were detected at comparable quantities in all tissues examined including spleens, livers and lungs between the experimental and control groups (Fig. 4d). Supporting these data, we did not observe any differences in terms of proliferation measured by BrdU incorporation as well as the number of cells undergoing apoptosis between WT and Atg7-deficient antigen specific CD8 T cells at day 5 p.i. when T cells are rapidly proliferating ( Supplementary Fig. 4d-h). An important function of CD8 T cells is the ability to produce the antiviral cytokines IFN-γ and TNF. Ex vivo stimulation of splenocytes at day 8 p.i. using peptides indicated little difference in the ability, or the number, of cells producing these cytokines (Fig. 4e). In addition, we no longer detected virus in the sera, spleen, liver or lung of wild type, nor Atg7 fl/fl Gzmb-Cre mice by day 8 p.i. (data not shown). Collectively, our data demonstrated that virus-specific CD8 T cells deficient in Atg7 were able to proliferate and differentiate into functional effector T cells that were comparable to their wild-type counterparts. This is consistent with our findings that autophagy is actually decreased in proliferating virus specific CD8 T cells (Figs. 1-3).
Immediately following the peak of expansion, the sharpest decline in virus-specific CD8 T cells was observed in the Atg7-deficient cells between days 8 and 15 p.i. (Fig. 4f-h). T cells specific for the dominant LCMV epitopes in Atg7 fl/fl Gzmb-Cre mice, including D b GP33, D b GP276, D b NP396 and K b GP34 were reduced approximately 5-10 fold in the spleens of Atg7 fl/fl Gzmb-Cre mice compared to wild type mice (Fig. 4g). Few antigen-specific T cells survived the contraction phase to transition into the memory phase (day 30 p.i.) within either lymphoid or non-lymphoid tissues, or the peripheral blood ( Fig. 4h and i). Overall, the data indicated that Atg7 is important for the survival of effector CD8 T cells, and further suggest that a defective autophagy pathway compromises the formation of a functional CD8 T cell memory pool. These results correlated very closely with the in vivo kinetics of autophagy. Thus, the striking survival defects of autophagy deficient effector T cells occur after day 8 p.i concurrently when autophagy is increased in virus specific T cells.
We sought to examine whether the ubiquitin-like conjugation systems necessary for autophagy were required for T cell memory formation by evaluating the importance of a second essential autophagy protein, Atg5. Similar to the Atg7 fl/fl Gzmb-Cre phenotype, Atg5 fl/fl Gzmb-Cre mice infected with LCMV Armstrong demonstrated an initial expansion (day 8 p.i.) followed by a drastic loss of antigen-specific CD8 T cells ( Fig. 5a and b). We observed this profound loss in antigen-specific T cells in all tissues examined at the memory phase in the Atg5 fl/fl Gzmb-Cre mice ( Fig. 5c and d). These data implied a pro-survival role for proteins involved in the autophagy membrane elongation complex in antigen-specific CD8 T cells. The conditional knockout system is a powerful tool to investigate functions of a gene within a specific cell population. However, one caveat of using this system is that the profound depletion of antigen-specific CD8 T cells observed in Atg7 fl/fl Gzmb-Cre and Atg5 fl/fl Gzmb-Cre mice during the course of LCMV infection (Figs. 4 and 5) may impact host environmental factors such as inflammatory cytokines that could exacerbate T cell survival and differentiation. Furthermore, other cell types, such as natural killer (NK) cells, also express granzyme B, introducing an additional level of complexity when comparing experimental and control mice. Recent work showed that NK cells were implicated in modulating T cell immunity 26,27 . To address these issues, we generated mixed bone marrow chimeras that were reconstituted from an mixture of congenitally marked Atg7 fl/fl Gzmb-Cre (CD45.2) and wild-type C57BL/6 (CD45.1) donor bone marrow cells into wild-type C57BL/6 (CD45.1) recipient mice ( Supplementary Fig. 5a). This system allowed us to compare changes in number between wild type and Atg7 fl/fl Gzmb-Cre antigen-specific CD8 T cells in the same environment (in the same mouse), therefore excluding factors that only act on one population of these cells. As a control, we also generated mixed bone marrow chimera mice that were reconstituted with Atg7 fl/fl (CD45.2) (no cre) and wild-type (CD45.1) donor mice. After reconstitution, these animals were challenged with the LCMV Armstrong, and virus-specific CD8 T cells were examined over time (Supplementary Fig.  5a). At day 8 p.i., there was a significant expansion of antigen-specific T cells from both CD45.2 (knock-out) and CD45.1 (wild type) donor populations. The D b GP33 and D b NP396-specific CD8 T cell responses were longitudinally analyzed during the effector and memory stages ( Fig. 6a and b, Supplementary Fig. 5c). Similar to the observation in the conditional knockout mice, we observed a more than 90% decline in Atg7-deficient LCMVspecific CD8 T cells (CD45.2 + ) from day 8 to day 15 in Atg7 fl/fl Gzmb-Cre plus C57BL/6 chimeric mice, compared to a moderate decline of 50%-60% in the corresponding wild-type (CD45.2 + ) population in the Atg7 fl/fl plus C57BL/6 controls. Four weeks after infection there were virtually no antigen-specific CD8 T cells of Atg7 fl/fl Gzmb-Cre origin (CD45.2 + ) throughout the peripheral blood, lymphoid and non-lymphoid tissues in Atg7 fl/fl Gzmb-Cre plus C57BL/6 chimeric mice (Fig. 6a-c). A significant proportion of naive CD8 T cells (CD44 lo ) from the Atg7 fl/fl Gzmb-Cre donor persisted within the chimeric mice, indicating specific loss of the antigen-experienced effector CD8 T cells from Atg7 fl/fl Gzmb-Cre mice throughout the progression of infection (Supplementary Fig. 5b). Since the cells from WT (CD45.1 + ) origin made up the majority of the reconstituted population within the chimeric mice ( Supplementary Fig. 5b), the physiological and pathological conditions closely resembled those of the WT mice. Hence, in the absence of potential external variables, such as changes introduced in other cell types or any subtle variations in viral clearance, the defects rising from Atg7-deficiency were cell-intrinsic properties that affected survival of the antigen-specific effector CD8 T cells to the memory phase. Thus, these data confirmed that the autophagy pathway is critical for memory T cell formation.

Distinct metabolic profiles of Atg7 −/− effector T cells
To investigate the potential mechanism underlying the survival defects in the Atg7-deficient CD8 T cells, we performed metabolomic and transcriptomic analyses on antigen-specific T cells from Atg7-deficient (Atg7 fl/fl Gzmb-Cre) or WT (Atg7 fl/fl ) mice 8 days following LCMV Armstrong infection. This time point was selected as representative of a transition time point in T cell differentiation before significant loss of Atg7-deficient cells has occurred. In the D b GP33 + CD8 T cells isolated from mice 8 day p.i, the overall gene expression pattern of the Atg7-deficient CD8 T cells closely resembled that of the WT group; only a small number of genes showed significantly different expression between the two groups in the DNA microarray data, mostly associated with cell cycle (data not shown). However, these cells exhibited distinct metabolic profiles measured on a liquid chromatography coupled mass spectrometry (LC-MS) metabolomics platform. Among the metabolites that were significantly different between Atg7-sufficent and deficient cells (Supplementary Table 1), many are contained within well-defined metabolic pathways with clear links to cell survival, as well as some of which have been implicated in T cell differentiation ( Supplementary Fig. 6). For example, among the pathways that were significantly different between genotypes are the carnitine shuttle and di-unsaturated fatty acid beta-oxidation, both of which are part of fatty acid metabolism in the mitochondria (Fig. 7a). T cell-intrinsic mitochondrial fatty acid oxidation has been shown to be a critical metabolic pathway for the stable development of long-lived memory T cells 16,17 . These data could indicate that autophagy contributes to the production of the lipid substrates for mitochondrial fatty acid oxidation and the fueling of oxidative phosphorylation (OXPHOS) in these cells 28 . Consistent with a role for autophagy in lipid metabolism in T cells during the transition to the memory phase, our metabolite analysis showed that lipid biosynthetic pathways were dysregulated in the absence of Atg7 (Fig. 7a). Also of note, glucosamine 6 phosphate and glycan biosynthesis pathways were perturbed in Atg7-deficient T cells ( Fig.  7a and Supplementary Fig. 6). The hexosamine pathway provides substrates for the glycosylation of growth factor cytokine receptors, leading to receptor stability and the maintenance of survival signals 29 . Given the importance of growth factor cytokines in supporting the development of memory T cells 2 , it is possible that T cells lacking autophagy machinery cannot maintain the proper growth factor signals that support memory T cell development, linking autophagy to glycosylation and the maintenance of cellular metabolism. Importantly, these metabolomics data were further validated by examining the levels of gene expression of the enzymes associated these metabolite activities. Using Gene set enrichment analysis 30 and these enzymes as a gene set, the gene expression for these metabolic activities was clearly associated with the Atg7 deficient genotype (Fig. 7b and Supplementary Table 2). Overall, several metabolic pathways were altered in Atg7-deficient T cells day 8 pi, and when integrated with our transcriptomic studies suggest that autophagy supports metabolic homeostasis in CD8 T cells during the transition to memory phase.

Autophagy in CD8 T cells is essential for viral control
Autophagy is required for memory CD8 T cell formation during an acute viral infection as shown above. Next, we asked whether autophagy regulates CD8 T cell responses during a chronic viral infection. To address this question, the Atg7 fl/fl Gzmb-Cre mice were challenged with the LCMV Clone 13 strain, which causes a chronic infection 31 . In chronic infections antigen-specific T cells gradually diminish in their effector functionality and become exhausted cells due to prolonged persistence of antigen in the host 32 . During a chronic infection, antigen-specific CD8 T cells in the Atg7 fl/fl Gzmb-Cre mice again mounted a initial response to the virus in the peripheral blood, as well as in lymphoid and nonlymphoid tissues at day 8 p.i., comparable to the wild-type controls (Fig. 8a, Supplementary  Fig. 7a and b). Ex vivo stimulation of splenocytes at day 8 p.i. showed little difference in the number of cells producing IFN-γ (Fig. 8b). However, by day 15 p.i., most LCMV-specific T cells were lost in Atg7 fl/fl Gzmb-Cre mice throughout lymphoid and non-lymphoid tissues, similar to the acute infection model (Fig. 8a, Supplementary Fig. 7c and d). The observed reduction was also reflected in the number of IFN-γ-producing cells when stimulated ex vivo (Fig. 8b). As a consequence of the loss of most antigen specific cells, viral control was compromised in Atg7 fl/fl Gzmb-Cre mice (Fig. 8c). These data further indicated a significant survival defect in the Atg7-deficient LCMV-specific effector CD8 T cells; not only were these cells compromised in memory CD8 T cell differentiation, but they also failed to survive in the chronic infection model.

DISCUSSION
Our findings that proliferating T cells show decreased autophagy in vivo were unexpected because previous in vitro studies showed that autophagy is increased in T cells after TCR stimulation 9,12,13 . Discrepancies between our current study and these previous studies may be explained by the vastly different condition of TCR stimulation in vivo versus in vitro. In addition, in some of the earlier studies only a single aspect of the autophagy pathway such as the number of autophagosomes and the amount of lipidated LC3b-II have been used to assess autophagy activity in cultured T cells. As the autophagy pathway can be blocked at any point along its progression, it has become clear that measuring a single component in the pathway, rather than measuring flux through the pathway as done here, is not reflective of whether autophagy is activated or blocked 7 .
Our findings of the kinetics of autophagy in vivo are totally consistent with our results in which essential autophagy genes were deleted in antigen specific CD8 T cells. The in vivo kinetics of autophagy indicate that activated T cells can proliferate with very low autophagy activity, and indeed both autophagy deficient and wild type virus-specific CD8 T cells were able to similarly proliferate in most tissues. However, this comparable proliferation of antigen specific CD8 T cells between autophagy gene conditional knockout and wild type mice was unexpected because a prior study showed that autophagy gene deletion resulted in compromised T cell proliferation after in vitro TCR stimulation 9 . T cells used in the prior study were autophagy defective for long periods of time before being assayed because autophagy genes were deleted during T cell development 9 . It is known that these autophagy gene deficient T cells are transcriptionally and phenotypically distinct from wild type T cells 9,10,15,33 . In contrast, to avoid this complication in our studies autophagy genes were specifically deleted in granzyme B expressing cells after T cell activation, and therefore naive T cells were normal at the time of viral infection. Thus, a robust CD8 T cell expansion in the absence of autophagy genes suggest that activated T cells can proliferate and differentiate into effectors without autophagy, and this notion is consistent with the in vivo kinetics of autophagy in antigen specific CD8 T cells.
In contrast to T cell expansion phase, autophagy was required for effector T cell survival to form memory T cells during contraction phase when effector T cells undergo significant changes. In this phase, effector T cells, having just finished many rounds of vigorous cell divisions, potentially accumulate protein aggregates, reactive oxygen species (ROS) and other superfluous cellular content 34,35,36,37,38 . Autophagy could perform multiple prosurvival functions in these cells, including the removal of damaged mitochondria that generate ROS or misfolded proteins 39,40 . Thus, upregulation of autophagy in virus-specific CD8 T cells upon conclusion of clonal expansion is important during the contraction phase.
Our results on the kinetics and the functional role of autophagy in virus specific CD8 T cells have important implications to previous studies linking the major regulator of autophagy, mTOR to the memory CD8 T cell differentiation 17,41 . These studies have demonstrated that inhibition of mTOR by rapamycin improves not only the magnitude but also quality of memory CD8 T cells 17,41 . These effects of rapamycin in antigen specific CD8 T cells might result from increased autophagy during the proliferation and contraction phase. Active mTOR signals directly down-regulate autophagy and inhibition of mTOR would, therefore, likely enhance autophagy activity 42 . Thus, it is an important future work to investigate whether rapamycin treatment up-regulates autophagy in antigen specific CD8 T cells and whether increased autophagy improves memory CD8 T cell differentiation.
Our metabolomics analysis provides insight into the metabolic pathways regulated by autophagy in virus-specific CD8 T cells. The unbiased metabolite data revealed that mitochondrial fatty acid oxidation, which has been shown to be critical for memory T cell generation 16,17 , was dysregulated in Atg7-deficient T cells at the peak of clonal expansion, right at the point Ag-specific T cells transition to the memory phase. A study of metabolomic profiles in a mouse cancer model reported that the Atg7-deficient tumor cells had defective mitochondria and reduced fatty acid oxidation 43 . We speculate that T cellintrinsic autophagy could be important for generating lipid substrates for mitochondrial fatty acid oxidation (FAO). This notion is consistent with published studies showing that autophagy can regulate lipid metabolism via the breakdown of cellular lipid stores 44 . Precisely how the substrates for mitochondrial FAO are utilized or generated by memory T cells, and whether autophagy directly regulates mitochondrial FAO in antigen specific effector CD8 T cells is an exciting subject for future work.
In addition to an acute viral infection, our data demonstrate that autophagy in virus-specific CD8 T cells is essential for cell survival and controlling a chronic viral infection. It is well established that activated CD8 T cells during chronic infections or in tumor environments differentiate into exhausted T cells. Rejuvenating these exhausted T cells is a promising approach to treat chronic infections and cancer. Thus, our data suggest that autophagy plays a critical role in formation and survival of exhausted CD8 T cells that can be targeted for immunotherapy.
In summary, these data establish that the autophagy pathway is dynamically regulated throughout the CD8 T cell response and is critical for the formation T cell memory. Autophagy seems to play a role in the metabolic homeostasis of effector CD8 T cells during their transition into memory cells. Our studies have implications to develop effective vaccination strategies and anti-cancer immune therapy through targeting the autophagy pathway and/or other metabolic pathways.

Mice and infection
Atg7-floxed mice and Atg5-floxed mice were generated and characterized previously 22,23 . Conditional knockout mice were generated by crossing Atg7-floxed mice and Atg5-floxed mice to Granzyme B-Cre transgenic mice 24 . To generate P14 chimeric mice bearing LCMV-specific CD8 T cells, transgenic P14 cells (Thy-1.1 + ) with an engineered TCR that recognizes the H-2D b GP33 epitope were adoptively transferred intravenously (1×10 4 antigen-specific CD8 + T cells per mouse as stated in the figure legend) to wild-type C57BL/6 mice (Jackson Laboratory). In acute infection, mice were infected with the Armstrong strain of lymphocytic choriomeningitis virus (LCMV) using an intraperitoneal (i.p.) injection of 2×10 5 plaque-forming unit (PFU). In chronic infection, mice were infected with the Clone 13 strain of LCMV using an intravenous injection of 2×10 6 PFU. All animal experiments were approved by the Institutional Animal Care and Use Committee of Emory University.

Cellular Imaging
Cells were analyzed on Image Stream Mark II (Amnis) using software INSPIRE. Images were acquired at 60x or 40x magnification with a low speed and high sensitivity setting. Single focused cells were selected for image collection. Data were analyzed using software IDEA. Spot counting analysis was used to evaluate LC3b puncta positive cells in each sample.

Cell sorting
Total activated CD8 T cells on day 8 post LCMV infection were stained with antibodies against CD8, CD44 and CD62L and were sorted using FACSAria II (BD). D b GP33-specific cells were stained with antibodies against CD8, CD44 as well as TCR using D b GP33 tetramer 46 . The typical purity after cell sorting was between 95-99%.

LC/MS metabolomic analysis
The metabolomics analysis was performed similarly as previously described (Li2013). Cell extracts were treated with acetonitrile (2:1, v/v) and centrifuged at 14,000× g for 5 min at 4°C to remove proteins. Samples were maintained at 4°C in an autosampler until injection. A Thermo Orbitrap-Velos mass spectrometer (Thermo Fisher, San Diego, CA) coupled with switching dual chromatography (an anion exchange (AE) column and a C18 reverse-phage column) was used for data collection, via positive-ion electrospray ionization (ESI). Each sample was run in 3 to 6 analytical replicates, depending on available source material. Mouse naive CD8 T cells, kidney and splenocyte tissues were also run as analytical controls. Metabolite features were extracted and quantified using apLCMS software version 5.9.8 47 .
Only features that were detected in 12 or more injection samples (including analytical replicates) were retained for further analysis. This filtering led to 3270 metabolite features from the AE column and 6300 from the C18 column. Data were log2 transformed, averaged among analytical replicates, and normalized by total ion intensity. Missing values were imputed with half mean of the feature in all detected samples. Student t-test was used to compare between KO and WT groups. Metabolic pathway analysis was performed by mummichog software 0.9.6 using default parameters 48 . Top 500 metabolite features in the ttest were used as input to mummichog, and the total list of features were used as reference.
Mummichog tested the enrichment of input metabolites against random data resampled from the reference list, and produced an empirical p-value per pathway 48 . Input metabolites in the significant pathways (p < 0.05 in both experiments) were linked in a network figure by known metabolic reactions (Supplementary Fig. 6).

Microarray analysis
Day 8 DbGP33-specific CD8 T cells from acute LCMV infected mice were FACS sorted based on CD8 and the tetramer stain. RNA was isolated using RNeasy kit (Qiangen) according to manufacturer's protocol. RNA was amplified, biotinylated and hydridized on mouse 430.2 Affymetrix microarray chips at the Microarray Core (Dana Farber Cancer Institute, MA, USA). The expression of metabolic enzymes underlying Supplementary Fig.  6 was tested for their association with KO or WT genotype. All enzyme commission (EC) numbers in the reactions in Supplementary Fig. 6 were looked up in mouse genome for their encoding genes. Only genes that are expressed in T cells are included for this analysis. The positive expression in T cells was according to the definition of ImmGen consortium, as signal intensity higher than 128 unit in any of their microarray files (total 89 microarrays on T cell populations, retrieved on Feb. 22 30 . Preranked method in GSEA was used, where the genes were ranked by t-statistic between KO and WT samples.

Statistical analysis
Statistical analysis was performed using GraphPad Prism 5. P values were calculated based on two-tailed unpaired Student's t-test, assuming normal distribution of data and equal sample variance. Sample size was not predetermined via statistical means, but the number of mice used in the study was consistent with previously published work of similar nature. All mice used for the study were included for a robust statistical analysis with no randomization or blinding involved.

Supplementary Material
Refer to Web version on PubMed Central for supplementary material.     Antigen-specific CD8 T cells lacking the Atg7 gene exhibit cell-intrinsic defects in developing into long-term memory cells in bone marrow chimeric mice. Atg7 fl/fl plus C57BL/6 (CD45.2/CD45.1) control and Atg7 fl/fl Gzmb-Cre plus C57BL/6 (CD45.2/CD45.1) experimental mixed bone marrow chimera mice were generated. CD8 T cell response was evaluated following the LCMV Armstrong infection according to the scheme in Supplementary Fig. 5a. (a)   Metabolomic and transcriptomic analysis of Atg7-dificient CD8 T cells. Atg7 fl/fl Gzmb-Cre (KO) and control (Atg7 fl/fl , WT) mice were infected with LCMV Armstrong. D b GP33 + CD8 T cells were isolated from mice 8 day p.i for analysis. (a) Pathways differentially regulated in Day 8 Atg7-deficient antigen-specific cells. Each column represents an independent experiment of samples pooled from 3 WT or KO mice; each row a pathway colored to indicate its p-value. (b) Gene set enrichment analysis (GSEA) of genes represented in T cells, which are associated with the metabolic enzymes underlying the most differentially regulated metabolic pathways (a) in KO vs. WT. p=0.024.