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| <?xml version="1.0" encoding="UTF-8"?><!DOCTYPE article PUBLIC "-//NLM//DTD JATS (Z39.96) Journal Archiving and Interchange DTD v1.1d1 20130915//EN" "JATS-archivearticle1.dtd"><article article-type="research-article" dtd-version="1.1d1" xmlns:xlink="http://www.w3.org/1999/xlink"><front><journal-meta><journal-id journal-id-type="nlm-ta">elife</journal-id><journal-id journal-id-type="hwp">eLife</journal-id><journal-id journal-id-type="publisher-id">eLife</journal-id><journal-title-group><journal-title>eLife</journal-title></journal-title-group><issn publication-format="electronic">2050-084X</issn><publisher><publisher-name>eLife Sciences Publications, Ltd</publisher-name></publisher></journal-meta><article-meta><article-id pub-id-type="publisher-id">03706</article-id><article-id pub-id-type="doi">10.7554/eLife.03706</article-id><article-categories><subj-group subj-group-type="display-channel"><subject>Research article</subject></subj-group><subj-group subj-group-type="heading"><subject>Immunology</subject></subj-group></article-categories><title-group><article-title>Autophagy is a critical regulator of memory CD8<sup>+</sup> T cell formation</article-title></title-group><contrib-group><contrib contrib-type="author" corresp="yes" id="author-15567"><name><surname>Puleston</surname><given-names>Daniel J</given-names></name><xref ref-type="aff" rid="aff1"/><xref ref-type="corresp" rid="cor1">*</xref><xref ref-type="other" rid="par-1"/><xref ref-type="other" rid="par-2"/><xref ref-type="other" rid="par-3"/><xref ref-type="fn" rid="con1"/><xref ref-type="fn" rid="conf1"/></contrib><contrib contrib-type="author" equal-contrib="yes" id="author-15608"><name><surname>Zhang</surname><given-names>Hanlin</given-names></name><xref ref-type="aff" rid="aff1"/><xref ref-type="fn" rid="equal-contrib">†</xref><xref ref-type="other" rid="par-1"/><xref ref-type="fn" rid="con2"/><xref ref-type="fn" rid="conf1"/></contrib><contrib contrib-type="author" equal-contrib="yes" id="author-15609"><name><surname>Powell</surname><given-names>Timothy J</given-names></name><xref ref-type="aff" rid="aff1"/><xref ref-type="fn" rid="equal-contrib">†</xref><xref ref-type="other" rid="par-1"/><xref ref-type="fn" rid="con6"/><xref ref-type="fn" rid="conf1"/></contrib><contrib contrib-type="author" id="author-15610"><name><surname>Lipina</surname><given-names>Elina</given-names></name><xref ref-type="aff" rid="aff1"/><xref ref-type="other" rid="par-1"/><xref ref-type="fn" rid="con3"/><xref ref-type="fn" rid="conf1"/></contrib><contrib contrib-type="author" id="author-12957"><name><surname>Sims</surname><given-names>Stuart</given-names></name><xref ref-type="aff" rid="aff2"/><xref ref-type="fn" rid="con7"/><xref ref-type="fn" rid="conf1"/></contrib><contrib contrib-type="author" id="author-19513"><name><surname>Panse</surname><given-names>Isabel</given-names></name><xref ref-type="aff" rid="aff1"/><xref ref-type="fn" rid="con4"/><xref ref-type="fn" rid="conf1"/></contrib><contrib contrib-type="author" id="author-15612"><name><surname>Watson</surname><given-names>Alexander S</given-names></name><xref ref-type="aff" rid="aff1"/><xref ref-type="other" rid="par-1"/><xref ref-type="other" rid="par-4"/><xref ref-type="other" rid="par-5"/><xref ref-type="fn" rid="con5"/><xref ref-type="fn" rid="conf1"/></contrib><contrib contrib-type="author" id="author-7889"><name><surname>Cerundolo</surname><given-names>Vincenzo</given-names></name><xref ref-type="aff" rid="aff1"/><xref ref-type="other" rid="par-1"/><xref ref-type="fn" rid="con8"/><xref ref-type="fn" rid="conf1"/></contrib><contrib contrib-type="author" equal-contrib="yes" id="author-15614"><name><surname>Townsend</surname><given-names>Alain RM</given-names></name><xref ref-type="aff" rid="aff1"/><xref ref-type="fn" rid="equal-contrib1">‡</xref><xref ref-type="other" rid="par-1"/><xref ref-type="fn" rid="con9"/><xref ref-type="fn" rid="conf1"/></contrib><contrib contrib-type="author" equal-contrib="yes" id="author-15615"><name><surname>Klenerman</surname><given-names>Paul</given-names></name><xref ref-type="aff" rid="aff2"/><xref ref-type="fn" rid="equal-contrib1">‡</xref><xref ref-type="other" rid="par-2"/><xref ref-type="fn" rid="con10"/><xref ref-type="fn" rid="conf1"/></contrib><contrib contrib-type="author" corresp="yes" id="author-9849"><name><surname>Simon</surname><given-names>Anna Katharina</given-names></name><xref ref-type="aff" rid="aff1"/><xref ref-type="corresp" rid="cor2">*</xref><xref ref-type="fn" rid="con11"/><xref ref-type="fn" rid="conf1"/></contrib><aff id="aff1"><institution content-type="dept">MRC Human Immunology Unit, Weatherall Institute of Molecular Medicine</institution>, <institution>University of Oxford, John Radcliffe Hospital</institution>, <addr-line><named-content content-type="city">Oxford</named-content></addr-line>, <country>United Kingdom</country></aff><aff id="aff2"><institution content-type="dept">Peter Medawar Building for Pathogen Research</institution>, <institution>University of Oxford</institution>, <addr-line><named-content content-type="city">Oxford</named-content></addr-line>, <country>United Kingdom</country></aff></contrib-group><contrib-group content-type="section"><contrib contrib-type="editor"><name><surname>Levine</surname><given-names>Beth</given-names></name><role>Reviewing editor</role><aff><institution>UT Southwestern Medical School</institution>, <country>United States</country></aff></contrib></contrib-group><author-notes><corresp id="cor1"><label>*</label>For correspondence: <email>daniel.puleston@ndm.ox.ac.uk</email> (DJP);</corresp><corresp id="cor2"><label>*</label>For correspondence: <email>katja.simon@ndm.ox.ac.uk</email> (AKS)</corresp><fn fn-type="con" id="equal-contrib"><label>†</label><p>These authors contributed equally to this work</p></fn><fn fn-type="con" id="equal-contrib1"><label>‡</label><p>These authors also contributed equally to this work</p></fn></author-notes><pub-date date-type="pub" publication-format="electronic"><day>11</day><month>11</month><year>2014</year></pub-date><pub-date pub-type="collection"><year>2014</year></pub-date><volume>3</volume><elocation-id>e03706</elocation-id><history><date date-type="received"><day>18</day><month>06</month><year>2014</year></date><date date-type="accepted"><day>16</day><month>10</month><year>2014</year></date></history><permissions><copyright-statement>© 2014, Puleston et al</copyright-statement><copyright-year>2014</copyright-year><copyright-holder>Puleston et al</copyright-holder><license xlink:href="http://creativecommons.org/licenses/by/4.0/"><license-p>This article is distributed under the terms of the <ext-link ext-link-type="uri" xlink:href="http://creativecommons.org/licenses/by/4.0/">Creative Commons Attribution License</ext-link>, which permits unrestricted use and redistribution provided that the original author and source are credited.</license-p></license></permissions><self-uri content-type="pdf" xlink:href="elife03706.pdf"/><abstract><object-id pub-id-type="doi">10.7554/eLife.03706.001</object-id><p>During infection, CD8<sup>+</sup> T cells initially expand then contract, leaving a small memory pool providing long lasting immunity. While it has been described that CD8<sup>+</sup> T cell memory formation becomes defective in old age, the cellular mechanism is largely unknown. Autophagy is a major cellular lysosomal degradation pathway of bulk material, and levels are known to fall with age. In this study, we describe a novel role for autophagy in CD8<sup>+</sup> T cell memory formation. Mice lacking the autophagy gene <italic>Atg7</italic> in T cells failed to establish CD8<sup>+</sup> T cell memory to influenza and MCMV infection. Interestingly, autophagy levels were diminished in CD8<sup>+</sup> T cells from aged mice. We could rejuvenate CD8<sup>+</sup> T cell responses in elderly mice in an autophagy dependent manner using the compound spermidine. This study reveals a cell intrinsic explanation for poor CD8<sup>+</sup> T cell memory in the elderly and potentially offers novel immune modulators to improve aged immunity.</p><p><bold>DOI:</bold> <ext-link ext-link-type="doi" xlink:href="10.7554/eLife.03706.001">http://dx.doi.org/10.7554/eLife.03706.001</ext-link></p></abstract><abstract abstract-type="executive-summary"><object-id pub-id-type="doi">10.7554/eLife.03706.002</object-id><title>eLife digest</title><p>In the face of an infection, the immune system mounts an aggressive response by producing many copies of killer immune cells called CD8<sup>+</sup> T cells that recognize and destroy any cells infected with the invading pathogen. The number of killer cells produced depends on the extent of the infection. Once the infection has been brought under control, most of the CD8<sup>+</sup> T cells die off. The small numbers that are retained—called memory cells—‘remember’ the pathogen, so that if it invades the body again, they can help the immune system to respond more quickly and effectively.</p><p>Memory cells are also critical to the effectiveness of vaccines, many of which introduce a dead or weakened pathogen into the body. This does not cause an infection, but does allow the immune system to create memory cells that are able to fend off the same pathogen in the future. However, vaccines only work in individuals that are able to produce and maintain memory cells, which many older people are less able to do.</p><p>An important system that maintains cells, called autophagy, destroys and removes the ‘junk’ and toxic by products that all cells accumulate over time as a result of normal cell functions. Without autophagy, cells become less able to produce energy and they may die. Puleston et al. show that autophagy begins to fail in old mice, which prevents the formation of a proper memory response. In addition, mice that lack an important gene needed for autophagy are unable to produce memory cells after being infected with viruses such as influenza.</p><p>Puleston et al. found that boosting autophagy in older mice using a chemical called spermidine—which is also found naturally in many tissues—helped to restore the mice's ability to create and maintain memory cells. Spermidine-treated mice developed a stronger immunity to influenza after vaccination compared with other mice of a similar age. Further research is required to better understand how spermidine works to see if it could be developed into a drug that safely boosts the immune system of humans.</p><p><bold>DOI:</bold> <ext-link ext-link-type="doi" xlink:href="10.7554/eLife.03706.002">http://dx.doi.org/10.7554/eLife.03706.002</ext-link></p></abstract><kwd-group kwd-group-type="author-keywords"><title>Author keywords</title><kwd>autophagy</kwd><kwd>memory T cells</kwd><kwd>vaccination</kwd><kwd>ageing</kwd><kwd>spermidine</kwd><kwd>influenza</kwd></kwd-group><kwd-group kwd-group-type="research-organism"><title>Research organism</title><kwd>mouse</kwd></kwd-group><funding-group><award-group id="par-1"><funding-source><institution-wrap><institution-id institution-id-type="FundRef">http://dx.doi.org/10.13039/501100000265</institution-id><institution>Medical Research Council</institution></institution-wrap></funding-source><award-id>Human Immunology Unit, Oxford</award-id><principal-award-recipient><name><surname>Puleston</surname><given-names>Daniel J</given-names></name><name><surname>Zhang</surname><given-names>Hanlin</given-names></name><name><surname>Powell</surname><given-names>Timothy J</given-names></name><name><surname>Lipina</surname><given-names>Elina</given-names></name><name><surname>Watson</surname><given-names>Alexander S</given-names></name><name><surname>Cerundolo</surname><given-names>Vincenzo</given-names></name><name><surname>Townsend</surname><given-names>Alain RM</given-names></name></principal-award-recipient></award-group><award-group id="par-2"><funding-source><institution-wrap><institution-id institution-id-type="FundRef">http://dx.doi.org/10.13039/100004440</institution-id><institution>Wellcome Trust</institution></institution-wrap></funding-source><award-id>088098/Z/08/Z</award-id><principal-award-recipient><name><surname>Puleston</surname><given-names>Daniel J</given-names></name><name><surname>Klenerman</surname><given-names>Paul</given-names></name></principal-award-recipient></award-group><award-group id="par-3"><funding-source><institution-wrap><institution>Allan and Nesta Ferguson Charitable Trust</institution></institution-wrap></funding-source><principal-award-recipient><name><surname>Puleston</surname><given-names>Daniel J</given-names></name></principal-award-recipient></award-group><award-group id="par-4"><funding-source><institution-wrap><institution-id institution-id-type="FundRef">http://dx.doi.org/10.13039/501100000038</institution-id><institution>Natural Sciences and Engineering Research Council of Canada</institution></institution-wrap></funding-source><principal-award-recipient><name><surname>Watson</surname><given-names>Alexander S</given-names></name></principal-award-recipient></award-group><award-group id="par-5"><funding-source><institution-wrap><institution-id institution-id-type="FundRef">http://dx.doi.org/10.13039/501100004101</institution-id><institution>Lady Tata Memorial Trust</institution></institution-wrap></funding-source><principal-award-recipient><name><surname>Watson</surname><given-names>Alexander S</given-names></name></principal-award-recipient></award-group><funding-statement>The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.</funding-statement></funding-group><custom-meta-group><custom-meta><meta-name>elife-xml-version</meta-name><meta-value>2</meta-value></custom-meta><custom-meta specific-use="meta-only"><meta-name>Author impact statement</meta-name><meta-value>Autophagy is required for CD8<sup>+</sup> T cell memory formation; in aged individuals autophagy levels are diminished, and increasing autophagy enhances their CD8<sup>+</sup> T cell responses.</meta-value></custom-meta></custom-meta-group></article-meta></front><body><sec id="s1" sec-type="intro"><title>Introduction</title><p>Upon successful clearance of a pathogen, the majority of short-lived effector T cells die and the remaining cells differentiate into a memory T cell population that provides long lasting immunity. While the cytokines, surface molecules, and signaling components involved in T cell memory formation have been extensively studied, the molecular pathways supporting these cell fate decisions are poorly understood. The T memory population is (1) quiescent, (2) long-lived and actively maintained, and (3) relies on mitochondrial respiration (<xref ref-type="bibr" rid="bib54">Pearce and Pearce, 2013</xref>). Both reduced cell cycling and longevity of this stem cell-like population demand rigorous maintenance of the cytoplasm as debris cannot be diluted to daughter cells, reminiscent of true stem cells (<xref ref-type="bibr" rid="bib55">Mortensen et al., 2011</xref>; <xref ref-type="bibr" rid="bib10">Guan et al., 2013</xref>; <xref ref-type="bibr" rid="bib49">Warr et al., 2013</xref>). Across mammalian cell types, clearance of debris and damaged organelles such as mitochondria is typically executed via autophagy (<xref ref-type="bibr" rid="bib7">Choi et al., 2013</xref>).</p><p>A recent study showed that formation of the influenza-specific B cell memory pool requires autophagy (<xref ref-type="bibr" rid="bib6">Chen et al., 2014</xref>). However, while the role of autophagy in the homeostasis of naïve T cells is well studied (<xref ref-type="bibr" rid="bib39">Puleston and Simon, 2014</xref>), nothing is known about the requirement of autophagy in antigen-experienced T cells. The T cell memory pool has previously been shown to be controlled by PI3K/Akt and AMPK signaling as well as mTOR (mammalian Target Of Rapamycin) inhibition (<xref ref-type="bibr" rid="bib3">Araki et al., 2009</xref>; <xref ref-type="bibr" rid="bib20">Kim et al., 2012</xref>; <xref ref-type="bibr" rid="bib42">Rolf et al., 2013</xref>), all of which also control autophagy (<xref ref-type="bibr" rid="bib17">Jung et al., 2010</xref>). Indeed, T cell memory responses can be improved with the mTOR inhibitor rapamycin (<xref ref-type="bibr" rid="bib3">Araki et al., 2009</xref>). Interestingly, rapamycin also rejuvenates HSCs in old mice (<xref ref-type="bibr" rid="bib5">Chen et al., 2009</xref>). Neither of these studies addressed autophagy as a contributing mechanism.</p><p>Evidence for age-related declining levels of autophagy stems largely from lower organisms such as yeast, flies, and worms (<xref ref-type="bibr" rid="bib43">Rubinsztein et al., 2011</xref>). A recent study showed that the polyamine spermidine induces autophagy and thereby prolongs life span in model organisms (<xref ref-type="bibr" rid="bib8">Eisenberg et al., 2009</xref>). Our own work demonstrated decreased levels of autophagy in CD8<sup>+</sup> T cells of old individuals (<xref ref-type="bibr" rid="bib34">Phadwal et al., 2012</xref>).</p><p>In this study, investigating a mouse model lacking the essential autophagy gene <italic>Atg7</italic> specifically in T cells, we find peripheral T cell lymphopenia, leading to proliferation and an activated phenotype within the CD8<sup>+</sup> T cell compartment. While <italic>Atg7</italic><sup><italic>−/−</italic></sup> T cells respond normally during the early stages of live viral challenge, a severely compromised memory CD8<sup>+</sup> T cell compartment was found in response to influenza and murine cytomegalovirus (MCMV). Using bone marrow (BM) chimeras, we excluded that this is due the effects of lymphopenia; poor CD4<sup>+</sup> T cell help; exhaustion, or altered cytokine receptor expression. Moreover, autophagy was found to be highest in antigen-specific CD8<sup>+</sup> T cells when compared to naïve cells. Antigen-specific <italic>Atg7</italic><sup><italic>−/−</italic></sup> CD8<sup>+</sup> T cells also underwent more cell death at the time of memory formation, display compromised mitochondrial health, and increased expression of the glucose receptor GLUT1, a marker for glycolysis. Furthermore, recall CD8<sup>+</sup> T cell responses to repeat immunizations and vaccination protocols were greatly diminished. This being reminiscent of the human ageing immune system (<xref ref-type="bibr" rid="bib11">Haq and McElhaney, 2014</xref>), we confirmed reduced autophagy at the transcriptional and functional level in murine T cells from old mice. Importantly, we were able to restore the CD8<sup>+</sup> T cell memory response in old mice with the autophagy-inducing compound spermidine, but not in autophagy-deficient mice. Finally, we found that spermidine induces autophagy independently of mTOR in T cells. Enhancing autophagy in an mTOR-independent manner may provide a safe way to improve vaccine responses in the elderly.</p></sec><sec id="s2" sec-type="results"><title>Results</title><sec id="s2-1"><title>Autophagy controls T cell numbers in naïve T<italic>-Atg7</italic><sup><italic>−/−</italic></sup> mice</title><p><italic>Atg7flox/flox</italic> mice were bred with <italic>CD4-Cre</italic> mice to generate mice with defective autophagy in both CD4<sup>+</sup> and CD8<sup>+</sup> T lymphocytes (T<italic>-Atg7</italic><sup><italic>−/−</italic></sup>). Successful excision and thereby absence of <italic>Atg7</italic> mRNA and Atg7 protein was confirmed in purified T cells (<xref ref-type="fig" rid="fig1s1">Figure 1—figure supplement 1A and B</xref>, respectively). Using the imaging flow cytometer (ImageStream) to count LC3 puncta in CD4<sup>+</sup> and CD8<sup>+</sup> T cells (<xref ref-type="bibr" rid="bib34">Phadwal et al., 2012</xref>), we demonstrated that functional autophagy was significantly diminished in <italic>Atg7</italic><sup><italic>−/−</italic></sup> CD8<sup>+</sup> T cells (<xref ref-type="fig" rid="fig1s1">Figure 1—figure supplement 1C</xref> with examples of ImageStream images in right panel). In addition, using a classical technique to detect lipidated LC3, we confirmed that basal autophagy was diminished in the presence and absence of the autophagy flux inhibitor Bafilomycin A (<xref ref-type="fig" rid="fig1s1">Figure 1—figure supplement 1D</xref>).</p><p>Previous reports have noted a number of changes to the naïve CD8<sup>+</sup> T cell compartment in the absence of autophagy, with T cell lymphopenia, a consistent observation (<xref ref-type="bibr" rid="bib36">Pua et al., 2007</xref>; <xref ref-type="bibr" rid="bib39">Puleston and Simon, 2014</xref>). We set out to investigate if an altered naïve CD8<sup>+</sup> T cell compartment exists in T<italic>-Atg7</italic><sup><italic>−/−</italic></sup> mice. We confirmed observations from previous reports using similar autophagy-deficient mouse models (<xref ref-type="bibr" rid="bib36">Pua et al., 2007</xref>, <xref ref-type="bibr" rid="bib37">2009</xref>) that thymic development of CD4<sup>+</sup> and CD8<sup>+</sup> T cells was normal in 6-week old T<italic>-Atg7</italic><sup><italic>−/−</italic></sup> mice (<xref ref-type="fig" rid="fig1">Figure 1A</xref>). However, mice were lymphopenic for both CD4<sup>+</sup> and CD8<sup>+</sup> T cells in the lymph nodes and blood (<xref ref-type="fig" rid="fig1">Figure 1B,C</xref>). Moreover, <italic>Atg7</italic><sup><italic>−/−</italic></sup> CD8<sup>+</sup> T cells exhibited an activated phenotype with increased CD44 expression (<xref ref-type="fig" rid="fig1">Figure 1D</xref>) and decreased CD62L expression (<xref ref-type="fig" rid="fig1">Figure 1E</xref>), resembling a ‘virtual memory’ compartment (<xref ref-type="bibr" rid="bib1">Akue et al., 2012</xref>). We observed similar frequencies of central effector memory CD62L<sup>+</sup>CD44<sup>hi</sup>, however, T-<italic>Atg7</italic><sup>−/−</sup> mice accumulated CD8<sup>+</sup> T cells with an effector memory phenotype (CD62L<sup>−</sup>CD44<sup>hi</sup>) (<xref ref-type="fig" rid="fig1s1">Figure 1—figure supplement 1E</xref>). Similarly, data from a mouse model in which another essential autophagy gene, <italic>Atg5</italic>, was deleted under the hematopoietic stem cell-specific promoter Vav, showed T cell lymphopenia and the expanded virtual memory T cell compartment (CD8<sup>+</sup>CD44<sup>+</sup>) suggesting this phenotype is not <italic>Atg7</italic> specific (<xref ref-type="fig" rid="fig1s2">Figure 1—figure supplement 2A and B</xref>). Next, we established that proliferation was increased in the activated CD44<sup>hi</sup> CD8<sup>+</sup> T cell compartment by Ki-67 staining (<xref ref-type="fig" rid="fig1">Figure 1F</xref>). The observed activated phenotype and increased cell turnover in <italic>Atg7</italic><sup><italic>−/−</italic></sup> CD8<sup>+</sup> T cells are likely driven by homeostatic proliferation in an attempt to fill the depleted T cell niche. Indeed, the expression of the homeostatic proliferation marker CD24 (<xref ref-type="bibr" rid="bib25">Li et al., 2006</xref>) was found to be significantly increased on <italic>Atg7</italic><sup><italic>−/−</italic></sup> CD8<sup>+</sup> T cells (<xref ref-type="fig" rid="fig1">Figure 1G</xref>). To investigate whether lymphopenia drives this activated phenotype in the CD8<sup>+</sup> T cell compartment, we generated 1:1 mixed bone marrow (BM) chimeras from CD45.2<sup>+</sup> T<italic>-Atg7</italic><sup><italic>−/−</italic></sup> BM mixed with CD45.1<sup>+</sup> wild-type BM. Both BMs contributed equally to form the new hematological system (<xref ref-type="fig" rid="fig1s2">Figure 1—figure supplement 2C</xref>). <italic>Atg7</italic><sup><italic>−/−</italic></sup> CD8<sup>+</sup> T cells were still diminished even in the presence of a replete T cell niche, suggesting the survival defect previously described for autophagy-deficient T cells (<xref ref-type="bibr" rid="bib36">Pua et al., 2007</xref>; <xref ref-type="bibr" rid="bib31">Mortensen et al., 2010</xref>) is cell-intrinsic (<xref ref-type="fig" rid="fig1s2">Figure 1—figure supplement 2D</xref>). However, the activated <italic>Atg7</italic><sup><italic>−/−</italic></sup> CD8<sup>+</sup> T cell phenotype was no longer detected in BM chimeras as measured by the frequency of donor CD45.2<sup>+</sup> CD8<sup>+</sup> T cells found to be CD62L<sup>+</sup> (<xref ref-type="fig" rid="fig1">Figure 1H</xref>) and CD44<sup>hi</sup> (<xref ref-type="fig" rid="fig1">Figure 1I</xref>). These data indicate that the observed homeostatic proliferation and the change in surface phenotype of <italic>Atg7</italic><sup><italic>−/−</italic></sup> CD8<sup>+</sup> T cells are driven by lymphopenia and are not cell-intrinsic.<fig-group><fig id="fig1" position="float"><object-id pub-id-type="doi">10.7554/eLife.03706.003</object-id><label>Figure 1.</label><caption><title>Lymphopenia induces homeostatic proliferation and an activated CD8<sup>+</sup> T cell phenotype in T-<italic>Atg7</italic><sup><italic>−/−</italic></sup> mice.</title><p>(<bold>A</bold>) Frequency of mature single positive CD4<sup>+</sup> and CD8<sup>+</sup> T cells in thymi of 6 week old mice (n = 4), representative FACS plot of three independent experiments. (<bold>B</bold>) Flow cytometric analysis of CD4<sup>+</sup> and CD8<sup>+</sup> T cell frequencies in blood and lymph nodes of T-<italic>Atg7</italic><sup><italic>−/−</italic></sup> and wild-type mice (dot plots depict staining in blood). Quantitative analyses are representative of six independent experiments, *p < 0.05 (n = 4). (<bold>C</bold>) Absolute counts of CD4<sup>+</sup> and CD8<sup>+</sup> T cells in blood of T-<italic>Atg7</italic><sup>−/−</sup> and WT mice over time (n = 4). (<bold>D</bold>) Percentage of CD44<sup>hi</sup> cells in the splenic CD8<sup>+</sup> T cell compartment of T-<italic>Atg7</italic><sup>−/−</sup> and WT mice. Bar graphs depict the frequency of gated cells (representative of seven independent experiments), *p < 0.05 (n = 4). (<bold>E</bold>) Percentage of splenic CD8<sup>+</sup> T cells positive for CD62L in T-<italic>Atg7</italic><sup>−/−</sup> and WT mice. Data are representative of three independent experiments, *p < 0.05, (n = 4). (<bold>F</bold>) Percentage of CD44<sup>lo</sup> and CD44<sup>hi</sup> CD8<sup>+</sup> T cells expressing Ki-67 in the spleen of T-<italic>Atg7</italic><sup>−/−</sup> and WT mice. Bar graphs are representative of three independent experiments, *p < 0.05 (n = 4). (<bold>G</bold>) Frequency of splenic CD8<sup>+</sup> T cells expressing CD24 in T-<italic>Atg7</italic><sup>−/−</sup> and WT mice. Bar graph is representative of two independent experiments, *p < 0.05 (n = 4). (<bold>H</bold>) Percentage of donor-derived CD62L<sup>+</sup>CD8<sup>+</sup> T cells. Lethally irradiated CD45.1 hosts reconstituted with a 1:1 mix of T-<italic>Atg7</italic><sup>−/−</sup> or WT BM (both CD45.2) with CD45.1 wild-type BM. Controls were WT and T-<italic>Atg7</italic><sup>−/−</sup> mice; *p < 0.05 (n = 4). (<bold>I</bold>) Frequency of donor-derived CD45.2<sup>+</sup> CD8<sup>+</sup> T cells in the spleen that is CD44<sup>hi</sup>. Controls were normal WT and T-<italic>Atg7</italic><sup>−/−</sup> mice, *p < 0.05 (n = 4). All values are mean ± s.e.m, and all statistical analyses are Mann–Whitney U-tests.</p><p><bold>DOI:</bold> <ext-link ext-link-type="doi" xlink:href="10.7554/eLife.03706.003">http://dx.doi.org/10.7554/eLife.03706.003</ext-link></p></caption><graphic xlink:href="elife03706f001"/></fig><fig id="fig1s1" position="float" specific-use="child-fig"><object-id pub-id-type="doi">10.7554/eLife.03706.004</object-id><label>Figure 1—figure supplement 1.</label><caption><title><italic>Atg7</italic> is efficiently excised in CD8<sup>+</sup> T cells from T-Atg7<sup>−/−</sup> mice resulting in loss of functional autophagy.</title><p>(<bold>A</bold>) <italic>Atg7</italic> gene expression in <italic>Atg7</italic><sup><italic>+/+</italic></sup> and <italic>Atg7</italic><sup><italic>−/−</italic></sup> T cells. T cells were isolated from the spleens of WT and T-<italic>Atg7</italic><sup><italic>−/−</italic></sup> mice by flow cytometry. mRNA was extracted, and <italic>Atg7</italic> gene expression was measured by q-PCR. Bar graph shows relative <italic>Atg7</italic> expression in T cells from T-<italic>Atg7</italic><sup><italic>−/−</italic></sup> mice compared to T cells from WT mice. ****p < 0.0001 by Student's <italic>t</italic> test. (<bold>B</bold>) Protein was purified from pooled MACS-sorted CD8<sup>+</sup> T cells from WT and T-<italic>Atg7</italic><sup><italic>−/−</italic></sup> mice and western blotted for ATG7. GAPDH was used as a loading control (n = 6 pooled mice). (<bold>C</bold>) LC3 Spot count in WT and <italic>Atg7</italic><sup><italic>−/−</italic></sup> CD8<sup>+</sup> T cells. Splenocytes from WT and T-<italic>Atg7</italic><sup><italic>−/−</italic></sup> mice were untreated or treated for 2 hr with an autophagy flux inhibitor before staining for relevant cell surface markers and LC3-II. LC3 spot count was assessed using ImageStream software. Quantification is gated on CD8<sup>+</sup> T cells. Right panel depicts example LC3 spot count images (×60 magnification). *p = 0.0132, ***p = 0.0002 as determined by Student <italic>t</italic> test (n = 4) (<bold>D</bold>) Autophagy flux in WT and <italic>Atg7</italic><sup><italic>−/−</italic></sup> CD8<sup>+</sup> T cells. Purified CD8<sup>+</sup> T cells were untreated or treated with bafilomycin for 6 hr before whole protein extraction and western blot for LC3. (<bold>E</bold>) Percentage of CD44<sup>hi</sup> CD62L<sup>−</sup> and CD44<sup>hi</sup> CD62L<sup>+</sup> cells in the splenic CD8<sup>+</sup> T cell compartment of T-<italic>Atg7</italic><sup>−/−</sup> and WT mice. Representative FACS plots of three independent experiments are shown, ***p = 0.0003 as determined by Mann–Whitney U-test (n = 9).</p><p><bold>DOI:</bold> <ext-link ext-link-type="doi" xlink:href="10.7554/eLife.03706.004">http://dx.doi.org/10.7554/eLife.03706.004</ext-link></p></caption><graphic xlink:href="elife03706fs001"/></fig><fig id="fig1s2" position="float" specific-use="child-fig"><object-id pub-id-type="doi">10.7554/eLife.03706.005</object-id><label>Figure 1—figure supplement 2.</label><caption><title>The T cell phenotype of Vav-Atg5<sup>−/−</sup> mice is similar to that of T-Atg7<sup>−/−</sup> mice and the survival defect of Atg7<sup>−/−</sup> CD8<sup>+</sup> T cells is cell intrinsic.</title><p>(<bold>A</bold>) Flow cytometric analysis of CD4<sup>+</sup> and CD8<sup>+</sup> T cell frequencies in the spleen of WT and Vav-<italic>Atg5</italic><sup><italic>−/−</italic></sup> mice. Quantitative analyses are representative of two independent experiments, **p < 0.01 (n = 5). (<bold>B</bold>) Frequency of CD44<sup>hi</sup> cells within the CD8<sup>+</sup> T cell subset of WT and Vav-<italic>Atg5</italic><sup><italic>−/−</italic></sup> mice. Data are representative of two independent experiments, **p < 0.01 (n = 5). (<bold>C</bold>) BM reconstitution in BM chimera mice. 9 weeks after marrow transplantation, the frequency of donor CD45.2 cells was assessed in the spleen by flow cytometry. Quantified is the frequency of splenocytes expressing CD45.2 in BM chimera mice and in normal WT and T-<italic>Atg7</italic><sup><italic>−/−</italic></sup> mice that acted as controls. (<bold>D</bold>) Frequency of CD8<sup>+</sup> T cells that are CD45.2 donor-derived in BM chimera mice. Example dot plots are gated on CD8<sup>+</sup> T cells. Quantified is the frequency of CD8<sup>+</sup> T cells that express CD45.2 in BM chimera and in normal WT and T-<italic>Atg7</italic><sup><italic>−/−</italic></sup> mice that acted as controls. *p < 0.05 (n = 4). Data are mean ± s.e.m and all statistical analyses are Mann–Whitney U-tests.</p><p><bold>DOI:</bold> <ext-link ext-link-type="doi" xlink:href="10.7554/eLife.03706.005">http://dx.doi.org/10.7554/eLife.03706.005</ext-link></p></caption><graphic xlink:href="elife03706fs002"/></fig></fig-group></p></sec><sec id="s2-2"><title>Autophagy controls the CD8<sup>+</sup> T cell memory pool</title><p>Previous studies have shown a role for autophagy in naive T cell organelle homeostasis and survival (<xref ref-type="bibr" rid="bib37">Pua et al., 2009</xref>; <xref ref-type="bibr" rid="bib16">Jia and He, 2011</xref>). However, the importance of autophagy in CD8<sup>+</sup> T cells responding to infection is unknown. To investigate this, we challenged <italic>T-Atg7</italic><sup><italic>−/−</italic></sup> mice with influenza (PR8 strain) and MCMV. In T<italic>-Atg7</italic><sup><italic>−/−</italic></sup> mice, we found normal expansion of the antigen-specific effector CD8<sup>+</sup> T cell (CD8<sup>+</sup> T<sub>eff</sub>) compartment using influenza-specific tetramers on day 10 (peak response) in the lungs (<xref ref-type="fig" rid="fig2">Figure 2A</xref>). However, due to the pre-existing lymphopenia observed in naïve mice, the absolute counts were diminished (<xref ref-type="fig" rid="fig2s1">Figure 2—figure supplement 1A</xref>). The CD8<sup>+</sup> T<sub>eff</sub> response to MCMV in the blood on day 7 was also normal (<xref ref-type="fig" rid="fig2">Figure 2B</xref>) (<xref ref-type="bibr" rid="bib14">Hutchinson et al., 2011</xref>). However, the ability of T<italic>-Atg7</italic><sup><italic>−/−</italic></sup> mice to form memory CD8<sup>+</sup> T cells (CD8<sup>+</sup> T<sub>mem</sub>) to both influenza and MCMV is severely compromised (<xref ref-type="fig" rid="fig2">Figure 2C,D</xref>). Performing serial bleeding on influenza infected mice over time, we demonstrated a catastrophic collapse of the antigen-specific CD8<sup>+</sup> T cell pool at day 21, resulting in a failure to retain CD8<sup>+</sup> T<sub>mem</sub> in T<italic>-Atg7</italic><sup><italic>−/−</italic></sup> mice (<xref ref-type="fig" rid="fig2">Figure 2E</xref>). Interestingly, the response to the ‘inflationary’ epitopes m38 and IE3 from MCMV is also compromised (<xref ref-type="fig" rid="fig2">Figure 2F</xref>). In response to these epitopes, wild-type CD8<sup>+</sup> T cells continue to expand throughout MCMV chronic infection. However, <italic>Atg7</italic><sup><italic>−/−</italic></sup> CD8<sup>+</sup> T cells fail to inflate and instead undergo a dramatic contraction leading to a failure to form a MCMV-specific CD8<sup>+</sup> T cell memory pool. This profound CD8<sup>+</sup> T cell contraction in T<italic>-Atg7</italic><sup><italic>−/−</italic></sup> mice is also observed in response to a conventional, non-inflating MCMV epitope (m45, <xref ref-type="fig" rid="fig2s1">Figure 2—figure supplement 1B</xref>). While viral titers against influenza in the lungs of wild-type and T-<italic>Atg7</italic><sup><italic>−/−</italic></sup> mice were comparable at day 3 of infection, they were significantly higher on day 6 in T-<italic>Atg7</italic><sup><italic>−/−</italic></sup> mice (<xref ref-type="fig" rid="fig2">Figure 2G</xref>). This is most likely due to the significantly lower absolute number of antigen-specific effector CD8<sup>+</sup> T cells. As T-<italic>Atg7</italic><sup><italic>−/−</italic></sup> mice survived influenza challenge, we expect that the virus is eventually cleared in all mice. In keeping with these findings, autophagy levels (CytoID) are significantly increased in the antigen-specific CD8<sup>+</sup> T cell compartment compared to naïve T cells (CD44<sup>lo</sup>) in response to influenza in both spleen (<xref ref-type="fig" rid="fig2">Figure 2H</xref>) and lungs (<xref ref-type="fig" rid="fig2">Figure 2I</xref>), indicating autophagy is induced upon antigen stimulation in vivo<italic>.</italic><fig-group><fig id="fig2" position="float"><object-id pub-id-type="doi">10.7554/eLife.03706.006</object-id><label>Figure 2.</label><caption><title>Normal effector CD8<sup>+</sup> T cell responses to viral infection but defective memory CD8<sup>+</sup> T cell formation in T-<italic>Atg7</italic><sup><italic>−/−</italic></sup> mice.</title><p>(<bold>A</bold>) Effector CD8<sup>+</sup> T cell response to influenza in WT and T-<italic>Atg7</italic><sup>−/−</sup> mice. Mice were immunized intra-nasally with 0.00032 HAU PR8 influenza. On day 10, antigen-specific CD8<sup>+</sup> T cells to nucleoprotein (NP) was assessed with tetramer in lungs. Dot plots show examples of tetramer staining gated on CD8<sup>+</sup> T cells. Bar graph indicates percentage of CD8<sup>+</sup> T cells specific for NP (n = 5–6) and is representative of three independent experiments. (<bold>B</bold>) Effector CD8<sup>+</sup> T cell's response to MCMV in WT and T-<italic>Atg7</italic><sup>−/−</sup> mice. CD8<sup>+</sup> T cells from blood were stained with m45 tetramer on day 7 post-infection. Dot plots show m45-tetramer<sup>+</sup> cells gated on CD8<sup>+</sup> T cells. Bar graph indicates % CD8<sup>+</sup> T cells specific for m45 (n = 4–5) and is representative of three independent experiments. (<bold>C</bold>) CD8<sup>+</sup> T<sub>mem</sub> response to influenza. WT and T-<italic>Atg7</italic><sup>−/−</sup> mice immunized as in (<bold>A</bold>) and the antigen-specific CD8<sup>+</sup> T cell response was assessed in lungs on day 50 by tetramer. *p < 0.05, by Mann–Whitney U-test (n = 4). Dot plots are gated on CD8<sup>+</sup> T cells. Bar graph is representative of three independent experiments. (<bold>D</bold>) CD8<sup>+</sup> T<sub>mem</sub>'s response to MCMV. Lung CD8<sup>+</sup> T cells on day 65 post-infection were stained with IE3-tetramer. Dot plots are gated on CD8<sup>+</sup> T cells. Quantitation depicts frequency of IE3 specific CD8<sup>+</sup> T cells. *p < 0.05, by Mann–Whitney U-test (n = 4). Data are representative of two independent experiments. (<bold>E</bold>) CD8<sup>+</sup> T cell kinetics to influenza infection. WT and T-<italic>Atg7</italic><sup>−/−</sup> mice were immunized as in (<bold>A</bold>) and CD8<sup>+</sup> T cell response tracked over time in blood by tetramer. Y-axis shows frequency of NP-specific CD8<sup>+</sup> T cells. (<bold>F</bold>) CD8<sup>+</sup> T cell kinetics to MCMV infection. CD8<sup>+</sup> T cell response to epitopes m38 (left panel) and IE3 (right panel) were tracked over time in blood by tetramer in WT and T-<italic>Atg7</italic><sup>−/−</sup> mice. Y-axis indicates the percentage of CD8<sup>+</sup> T cells that are m38-specific. (<bold>G</bold>) Influenza virus titres. WT and T-<italic>Atg7</italic><sup><italic>−/−</italic></sup> mice were culled at days 3 and 6 post-immunization with PR8, and lungs were collected and snap frozen in liquid nitrogen. Virus titres were determined using MDCK-SIAT1 cells. ***p = 0.0002 as determined by Student t-test (n = 4) (<bold>H</bold>) WT mice were immunized with PR8 influenza as in (<bold>A</bold>). CD8<sup>+</sup> T cells from spleen were stained with CytoID at day 9 post-infection and assessed by flow cytometry. Histograms show examples of CD44<sup>lo</sup> CD8<sup>+</sup> T cells from unimmunized mice (filled grey line) and in NP-specific CD8<sup>+</sup> T cells from immunized mice (open black line). Quantification is by mean fluorescence intensity (MFI) of CytoID on gated indicated cell population and representative of two independent experiments. ****p < 0.0001 by Student <italic>t</italic> test (n = 6). (<bold>I</bold>) Autophagy levels by CytoID staining on CD8<sup>+</sup> T cells from the lungs on day 9 post-infection as in (<bold>G</bold>). Histograms provide examples of CytoID staining, gated on CD8<sup>+</sup> T cells in lungs of immunized mice (solid grey lines), and gated on NP-tetramer-specific CD8<sup>+</sup> T cells from immunized mice (open black lines). Quantification is by mean fluorescence intensity (MFI) of CytoID on gated indicated cell population and representative of two independent experiments. ****p < 0.0001 by Student <italic>t</italic> test (n = 4–8). All values are mean ± s.e.m.</p><p><bold>DOI:</bold> <ext-link ext-link-type="doi" xlink:href="10.7554/eLife.03706.006">http://dx.doi.org/10.7554/eLife.03706.006</ext-link></p></caption><graphic xlink:href="elife03706f002"/></fig><fig id="fig2s1" position="float" specific-use="child-fig"><object-id pub-id-type="doi">10.7554/eLife.03706.007</object-id><label>Figure 2—figure supplement 1.</label><caption><title><italic>T-Atg7<sup>−/−</sup></italic> mice fail to form memory CD8<sup>+</sup> T cells to a conventional MCMV epitope.</title><p>(<bold>A</bold>) Effector CD8<sup>+</sup> T cell absolute counts in WT and T-<italic>Atg7</italic><sup>−/−</sup> mice. Quantification of the absolute number of CD8<sup>+</sup> NP-tetramer<sup>+</sup> T cells was determined in the lungs on day 10 of PR8 influenza infection. **p = 0.0084 by Student <italic>t</italic> test (n = 6–9). (<bold>B</bold>) Frequency of m45-specific CD8<sup>+</sup> T cells in the liver of MCMV-immunized WT and T-<italic>Atg7</italic><sup><italic>−/−</italic></sup> mice on day 100 post-infection. Dot plots are gated on CD8<sup>+</sup> T cells; bar graphs depict the percentage of CD8<sup>+</sup> T cells that are m45-tetramer<sup>+</sup>. All values are mean ± s.e.m.</p><p><bold>DOI:</bold> <ext-link ext-link-type="doi" xlink:href="10.7554/eLife.03706.007">http://dx.doi.org/10.7554/eLife.03706.007</ext-link></p></caption><graphic xlink:href="elife03706fs003"/></fig></fig-group></p></sec><sec id="s2-3"><title>The requirement for autophagy in memory CD8<sup>+</sup> T cell differentiation is cell intrinsic</title><p>To exclude that a failure to maintain a CD8<sup>+</sup> T<sub>mem</sub> pool is caused by lymphopenia in T<italic>-Atg7</italic><sup><italic>−/−</italic></sup> mice, we performed viral challenge experiments in 1:1 mixed BM chimeras, facilitating the observation of antigen-specific <italic>Atg7</italic><sup><italic>−/−</italic></sup> CD8<sup>+</sup> T cell responses in a replete T cell environment. Even in the context of a BM chimera, <italic>Atg7</italic><sup><italic>−/−</italic></sup> CD8<sup>+</sup> T cells fail to generate a CD8<sup>+</sup> T<sub>mem</sub> pool following influenza infection, suggesting this defect is cell intrinsic (<xref ref-type="fig" rid="fig3">Figure 3A</xref>). By using BM chimeras to observe antigen-specific <italic>Atg7</italic><sup><italic>−/−</italic></sup> CD8<sup>+</sup> T cell responses, we excluded a number of explanations as to why T-<italic>Atg7</italic><sup><italic>−/−</italic></sup> mice fail to form CD8<sup>+</sup> T<sub>mem</sub>: (1) lymphopenia, (2) defective CD4<sup>+</sup> T cell help, and (3) the excision of <italic>Atg7</italic> in CD4-expressing antigen-presenting cells affecting CD8<sup>+</sup> T cell priming. Furthermore, a defect in the frequency of memory precursor cells might also impact upon CD8<sup>+</sup> T<sub>mem</sub> formation in T<italic>-Atg7</italic><sup><italic>−/−</italic></sup> mice. Mixed BM chimeras demonstrated that the frequency of short-lived effector (SLEC, CD127<sup>+</sup>KLRG1<sup>−</sup>) and memory precursor (MPEC, CD127<sup>−</sup>KLRG1<sup>+</sup>) CD8<sup>+</sup> T cells was in fact normal in T<italic>-Atg7</italic><sup><italic>−/−</italic></sup> mice (<xref ref-type="fig" rid="fig3">Figure 3B</xref>). We then tested whether the lack of memory response is due to exhaustion of autophagy-deficient antigen-specific CD8<sup>+</sup> T cells. While in T<italic>-Atg7</italic><sup><italic>−/−</italic></sup> mice, we found a sharp increase of antigen-specific CD8<sup>+</sup> T cells co-expressing the exhaustion markers PD-1<sup>+</sup> and TIM-3<sup>+</sup>; this was not found in the mixed BM chimeras. This indicates that without lymphopenia, exhaustion is not a hallmark of <italic>Atg7</italic><sup><italic>−/−</italic></sup> CD8<sup>+</sup> T cell responses and cannot explain the diminished CD8<sup>+</sup> T<sub>mem</sub> compartment (<xref ref-type="fig" rid="fig3">Figure 3C</xref>). We then sought to determine if altered expression of cytokine receptors crucial for CD8<sup>+</sup> T<sub>mem</sub> maintenance could explain the defective CD8<sup>+</sup> T<sub>mem</sub> compartment. However, both Il-7Rα (CD127) and Il-15Rα were normally expressed on <italic>Atg7</italic><sup><italic>−/−</italic></sup> MCMV-specific CD8<sup>+</sup> T cells (<xref ref-type="fig" rid="fig3">Figure 3D,E</xref>).<fig-group><fig id="fig3" position="float"><object-id pub-id-type="doi">10.7554/eLife.03706.008</object-id><label>Figure 3.</label><caption><title>Loss of memory CD8<sup>+</sup> T cell formation in the absence of <italic>Atg7</italic> is not due to lymphopenia, poor CD4<sup>+</sup> T cell help, exhaustion, or defective cytokine receptor expression.</title><p>(<bold>A</bold>) CD8<sup>+</sup> T<sub>mem</sub> response to influenza in mixed BM chimeras, generated as in <xref ref-type="fig" rid="fig1">Figure 1H</xref>. Mice were immunized with PR8 influenza, and the CD8<sup>+</sup> T<sub>mem</sub> response of the CD45.2 donor to NP was assessed in lungs by tetramer on day 40. Quantitation shows frequency of (donor) CD45.2<sup>+</sup> CD8<sup>+</sup> T cells that are NP-specific (n = 5–6). **p < 0.01, by Mann–Whitney U-test (n = 4–7). (<bold>B</bold>) SLEC and MPEC populations in the <italic>Atg7</italic><sup><italic>+/+</italic></sup> and <italic>Atg7</italic><sup><italic>−/−</italic></sup> antigen-specific CD8<sup>+</sup> T cell pool. Mixed BM chimera generated as in <xref ref-type="fig" rid="fig1">Figure 1H</xref>, were immunized with MCMV 8 weeks after transplantation. Dot plots show example of KLRG1 and CD127 expression on gated CD45.2<sup>+</sup> m45-tetramer<sup>+</sup> CD8<sup>+</sup> T cells on day 10 post-infection. Upper bar graph depicts the % of CD45.2<sup>+</sup> m45-tetramer<sup>+</sup> CD8<sup>+</sup> T cells that are CD127<sup>−</sup> KLRG1<sup>+</sup> (SLECs). Lower bar graph shows the % of CD127<sup>+</sup> KLRG1<sup>−</sup> (MPECs) in the same population. *p < 0.05, by Mann–Whitney U-test (n = 4–7). (<bold>C</bold>) Markers of exhaustion on <italic>Atg7</italic><sup><italic>−/−</italic></sup> MCMV-specific CD8<sup>+</sup> T cells on MCMV challenged BM chimera. Dot plots depict example of PD-1 and TIM-3 staining on gated CD45.2<sup>+</sup> m45-tetramer<sup>+</sup> CD8<sup>+</sup> T cells. Bar graph quantifies the percentage of (donor) CD45.2<sup>+</sup> m45-tetramer<sup>+</sup> CD8<sup>+</sup> T cells that are PD-1<sup>+</sup> TIM-3<sup>+</sup> at day 10 post-infection. *p < 0.05, by Mann–Whitney U-test (n = 4–7). (<bold>D</bold>) CD127 expression on <italic>Atg7</italic><sup><italic>−/−</italic></sup> MCMV-specific CD8<sup>+</sup> T cells in MCMV challenged BM chimeras. Examples of CD127 staining on gated CD45.2<sup>+</sup> m45-tetramer<sup>+</sup> CD8<sup>+</sup> T cells from spleen on day 10 post-infection are shown. *p < 0.05, by Mann–Whitney U-test (n = 4–7). (<bold>E</bold>) IL-15Rα expression on splenic <italic>Atg7</italic><sup><italic>−/−</italic></sup> MCMV-specific CD8<sup>+</sup> T cells in MCMV challenged BM chimeras. Histograms depict IL-15Rα expression in CD44<sup>lo</sup> CD8<sup>+</sup> T cells from unimmunized mice (grey dotted line), <italic>Atg7</italic><sup><italic>+/+</italic></sup> CD45.2<sup>+</sup> m45-tetramer CD8<sup>+</sup> T cells (grey filled line), and <italic>Atg7</italic><sup><italic>−/−</italic></sup> CD45.2<sup>+</sup> m45-tetramer CD8<sup>+</sup> T cells (black line). The left histogram shows expression in control normal WT and T-<italic>Atg7</italic><sup><italic>−/−</italic></sup> mice, the right histogram indicates staining in donor CD45.2<sup>+</sup> cells from BM chimera mice. Quantified is IL-15Rα mean fluorescence intensity on gated CD45.2<sup>+</sup> m45-tetramer CD8<sup>+</sup> T cells (n = 4–7). All values are mean ± s.e.m.</p><p><bold>DOI:</bold> <ext-link ext-link-type="doi" xlink:href="10.7554/eLife.03706.008">http://dx.doi.org/10.7554/eLife.03706.008</ext-link></p></caption><graphic xlink:href="elife03706f003"/></fig><fig id="fig3s1" position="float" specific-use="child-fig"><object-id pub-id-type="doi">10.7554/eLife.03706.009</object-id><label>Figure 3—figure supplement 1.</label><caption><title>Failure of to form memory CD8<sup>+</sup> T cells in T-<italic>Atg7<sup>−/−</sup></italic> mice is not the result of defective IRF4 or EOMES expression.</title><p>(<bold>A</bold>) IRF4 expression in <italic>Atg7</italic><sup><italic>−/−</italic></sup> and <italic>Atg7</italic><sup><italic>+/+</italic></sup> splenic m45-specific CD8<sup>+</sup> T cells on day 9, 15, and 22 post-infection. As a control, IRF4 was also measured in CD44<sup>lo</sup> CD8<sup>+</sup> T cells from unimmunized mice (naïve). Quantification shows IRF4 mean fluorescence intensity from gated m45-tetramer<sup>+</sup> CD8<sup>+</sup> T cells and CD44<sup>lo</sup> CD8<sup>+</sup> T cells (naïve). Statistics—Student's <italic>t</italic> test (n = 4–5). (<bold>B</bold>) EOMES expression in <italic>Atg7</italic><sup><italic>−/−</italic></sup> and <italic>Atg7</italic><sup><italic>+/+</italic></sup> antigen-specific CD8<sup>+</sup> T cells. WT and T-<italic>Atg7</italic><sup><italic>−/−</italic></sup> mice were immunized with MCMV and EOMES expression was measured in m45-specific CD8<sup>+</sup> T cells on day 9, 15, and 22. As a control, EOMES was also measured in CD44<sup>lo</sup> CD8<sup>+</sup> T cells from unimmunized mice (naïve). Quantification shows EOMES mean fluorescence intensity on gated m45-tetramer<sup>+</sup> CD8<sup>+</sup> T cells and CD44<sup>lo</sup> CD8<sup>+</sup> T cells (naïve) (n = 4–5).</p><p><bold>DOI:</bold> <ext-link ext-link-type="doi" xlink:href="10.7554/eLife.03706.009">http://dx.doi.org/10.7554/eLife.03706.009</ext-link></p></caption><graphic xlink:href="elife03706fs004"/></fig></fig-group></p><p>Finally, we tested whether autophagy compromises effector/memory differentiation and measured two transcription factors typically required during this differentiation. We found that the expression of IRF4, responsible for sustaining the expansion and differentiation of CD8<sup>+</sup> T<sub>eff</sub> (<xref ref-type="bibr" rid="bib52">Yao et al., 2013</xref>), and EOMES, a factor that promotes CD8<sup>+</sup> T<sub>mem</sub> formation (<xref ref-type="bibr" rid="bib15">Intlekofer et al., 2005</xref>), behaved normally over time in <italic>Atg7</italic><sup><italic>−/−</italic></sup> CD8<sup>+</sup> T cells (<xref ref-type="fig" rid="fig3s1">Figure 3—figure supplement 1A and B</xref>).</p></sec><sec id="s2-4"><title>Autophagy is required to maintain healthy mitochondria and cell viability for antigen-specific CD8<sup>+</sup> T cells</title><p>To understand the mechanism that leads to the failure to maintain a memory compartment in the absence of autophagy, we next checked if <italic>Atg7</italic><sup><italic>−/−</italic></sup> CD8<sup>+</sup> T cells were undergoing more cell death than wild-type cells. While this is well described for naïve autophagy deficient T cells (<xref ref-type="bibr" rid="bib39">Puleston and Simon, 2014</xref>), it had not been analyzed in antigen-specific T cells. As expected, in wild-type mice challenged with MCMV, we found highest levels of cell death among antigen-specific CD8<sup>+</sup> T cells just after the peak of the effector phase on day 9, whereas in T-<italic>Atg7</italic><sup><italic>−/−</italic></sup> mice, cell death was found to increase over time (<xref ref-type="fig" rid="fig4">Figure 4A</xref>). This could not be explained by a change in levels of the anti-apoptotic protein Bcl-2 (<xref ref-type="fig" rid="fig4s1">Figure 4—figure supplement 1A</xref>). The control of mitochondrial quality and reactive oxygen species (ROS) via mitophagy, the degradation of mitochondria, has been found to prevent cell death in T cells (<xref ref-type="bibr" rid="bib37">Pua et al., 2009</xref>). In keeping with this, mitochondrial content (<xref ref-type="fig" rid="fig4">Figure 4B</xref>) and mitochondrial ROS (<xref ref-type="fig" rid="fig4">Figure 4C</xref>) were significantly increased in <italic>Atg7</italic><sup><italic>−/−</italic></sup> antigen-specific CD8<sup>+</sup> T cells. While this likely explains the increased cell death, as shown for other hematopoietic cells (<xref ref-type="bibr" rid="bib31">Mortensen et al., 2010</xref>) and T cells (<xref ref-type="bibr" rid="bib37">Pua et al., 2009</xref>), mitophagy is also thought to be essential for maintenance of healthy mitochondrial energy generation. This notion is consistent with our data in other <italic>autophagy-deficient</italic> hematopoietic cell types (macrophages and primary leukemic lines, submitted) demonstrating increased glycolytic enzymes by proteomic analysis, increased lactate production and decreased oxygen consumption using metabolic seahorse measurements. Indeed, studies by Pearce et al showed that formation of the CD8<sup>+</sup> T<sub>mem</sub> pool is accompanied by a switch to mitochondrial respiration (<xref ref-type="bibr" rid="bib33">Pearce et al., 2009</xref>; <xref ref-type="bibr" rid="bib46">Sukumar et al., 2013</xref>). However, in the physiological viral infection models used here, aiming to mimic human infection, the scarcity of antigen-specific CD8<sup>+</sup> T cells prevented us from performing these metabolic measurements. In addition, due to the loss of CD8<sup>+</sup> T<sub>mem</sub> in the absence of <italic>Atg7</italic> by day 30, any metabolic analysis was restricted to early time points when antigen-specific CD8<sup>+</sup> T cells are still present in T<italic>-Atg7</italic><sup><italic>−/−</italic></sup> mice. We resorted to the measurement of a surrogate marker of glycolysis, the glucose transporter GLUT-1 on antigen-specific CD8<sup>+</sup> T cells by two different techniques, (a) the fluorescently labeled GLUT-1 binding domain of HTLV (human T cell leukemia virus), which specifically detects GLUT-1 (<xref ref-type="bibr" rid="bib26">Manel et al., 2003</xref>; <xref ref-type="bibr" rid="bib21">Kinet et al., 2007</xref>), and (b) a GLUT-1 antibody staining. As expected, GLUT-1 is upregulated on CD8<sup>+</sup> T<sub>eff</sub> (day 9) and then downregulated on CD8<sup>+</sup> T<sub>mem</sub> (day 22) in wild-type mice as CD8<sup>+</sup> T cells switch to mitochondrial respiration (<xref ref-type="fig" rid="fig4">Figure 4D</xref>). However, antigen-specific <italic>Atg7</italic><sup><italic>−/−</italic></sup> CD8<sup>+</sup> T<sub>eff</sub> expresses more GLUT-1 and downregulation does not occur to the same extent at the later time points compared to WT cells (day 22) (<xref ref-type="fig" rid="fig4">Figure 4D</xref>). This suggests that autophagy supports the metabolic switch during the differentiation from T<sub>eff</sub> to T<sub>mem</sub>. This was confirmed using the GLUT-1 antibody (<xref ref-type="fig" rid="fig4s1">Figure 4—figure supplement 1B</xref>). However, treatment with the anti-diabetic drug metformin that induces mitochondrial β-oxidation metabolism in T cells (<xref ref-type="bibr" rid="bib33">Pearce et al., 2009</xref>) did not rescue the memory T cell compartment in this model (data not shown).<fig-group><fig id="fig4" position="float"><object-id pub-id-type="doi">10.7554/eLife.03706.010</object-id><label>Figure 4.</label><caption><title><italic>Atg7</italic><sup><italic>−/−</italic></sup> memory CD8<sup>+</sup> T cells show increased mitochondrial content, reactive oxygen species, apoptosis and fail to down-regulate GLUT-1.</title><p>(<bold>A</bold>) The spleens of unimmunized and MCMV-infected mice were stained with the apoptotic marker Annexin V and a dead cell dye that stains cells with disrupted membranes on the time points indicated. Apoptotic cells were defined as dead cell dye-negative Annexin V<sup>+</sup>. Dot plots are gated on either CD44<sup>lo</sup> CD8<sup>+</sup> T cells (naïve) or m45-tetramer<sup>+</sup> CD8<sup>+</sup> T cells. *p < 0.05, by Mann–Whitney U-test (n = 4–5). (<bold>B</bold>) Mitochondrial volume by MitoTracker Green in Tetramer<sup>+</sup> CD8<sup>+</sup> T cells from WT and T-<italic>Atg7</italic><sup>−/−</sup> mice. Spleens from MCMV-immunized mice were stained with MitoTracker Green on day 15 post-infection. Quantification depicts mean fluorescence intensity on m45-tetramer<sup>+</sup> CD8<sup>+</sup> T cells and is representative of three independent experiments. **p < 0.01, Student <italic>t</italic> test (n = 4–5). (<bold>C</bold>) Mitochondrial superoxide production in Tetramer<sup>+</sup> CD8<sup>+</sup> T cells by MitoSox. Spleens from MCMV-immunized WT and T-<italic>Atg7</italic><sup>−/−</sup> mice were stained with MitoSox Red on day 15 post-infection and analyzed by flow cytometry. Bar graph depicts mean fluorescence intensity on m45-tetramer<sup>+</sup> CD8<sup>+</sup> T cells and is representative of three independent experiments. **p < 0.01, by Student <italic>t</italic> test (n = 4–5). (<bold>D</bold>) GLUT-1 expression. Tetramer<sup>+</sup> CD8<sup>+</sup> T cells from MCMV-immunized WT and T-<italic>Atg7</italic><sup><italic>−/−</italic></sup> mice were stained with the GFP-tagged HTLV receptor binding domain (eGFP-H<sub>RBD</sub>), that binds GLUT-1, at the time points indicated. As a control, GLUT-1 was also measured on CD44<sup>lo</sup> CD8<sup>+</sup> T cells from unimmunized mice (naïve). Bar graph shows the percentage of cells expressing GLUT-1. *p < 0.05, as determined by Mann–Whitney U-test (n = 4–5). All values are mean ± s.e.m.</p><p><bold>DOI:</bold> <ext-link ext-link-type="doi" xlink:href="10.7554/eLife.03706.010">http://dx.doi.org/10.7554/eLife.03706.010</ext-link></p></caption><graphic xlink:href="elife03706f004"/></fig><fig id="fig4s1" position="float" specific-use="child-fig"><object-id pub-id-type="doi">10.7554/eLife.03706.011</object-id><label>Figure 4—figure supplement 1.</label><caption><title>Normal Bcl-2 levels and altered GLUT-1 expression on antigen-specific <italic>Atg7<sup>−/−</sup></italic> CD8<sup>+</sup> T cells.</title><p>(<bold>A</bold>) MCMV-immunized WT and T-<italic>Atg7</italic><sup><italic>−/−</italic></sup> mice were assessed for Bcl-2 expression in splenic m45-specific CD8<sup>+</sup> T cells on day 9 and 22 post-infection. As a control, Bcl-2 was measured in CD44<sup>lo</sup> CD8<sup>+</sup> T cells from unimmunized mice (naïve). Quantification shows Bcl-2 mean fluorescence intensity. Statistics—Student's <italic>t</italic> test (n = 4–5). (<bold>B</bold>) GLUT-1 antibody staining on day 9 and day 22 post-infection in MCMV-immunized WT and T-<italic>Atg7</italic><sup><italic>−/−</italic></sup> mice. As a control, GLUT-1 was assessed in CD44<sup>lo</sup> CD8<sup>+</sup> T cells from unimmunized mice (naïve). Bar graphs indicate the frequency of m45-tetramer<sup>+</sup> CD8<sup>+</sup> T cells and CD44<sup>lo</sup> CD8<sup>+</sup> T cells (naïve) that express GLUT-1. Statistics—Mann Whitney U-test (n = 4–5).</p><p><bold>DOI:</bold> <ext-link ext-link-type="doi" xlink:href="10.7554/eLife.03706.011">http://dx.doi.org/10.7554/eLife.03706.011</ext-link></p></caption><graphic xlink:href="elife03706fs005"/></fig></fig-group></p></sec><sec id="s2-5"><title>Autophagy deficiency leads to diminished recall responses</title><p>We then tested whether T<italic>-Atg7</italic><sup><italic>−/−</italic></sup> mice were able to mount a recall CD8<sup>+</sup> T cell response to a secondary infection. We primed T<italic>-Atg7</italic><sup><italic>−/−</italic></sup> mice with PR8 (H1N1) followed by heterologous challenge with the X31 strain of influenza (H3N2). By challenging with a heterologous virus strain expressing different surface antigens, it is possible to significantly diminish the influence of antibodies in mediating immunity to secondary infection. Thus, heterotypic immunity relies heavily on cross-reactive CD8<sup>+</sup> T cell responses (<xref ref-type="bibr" rid="bib53">Zweerink et al., 1977</xref>; <xref ref-type="bibr" rid="bib47">Townsend et al., 1986</xref>), as opposed to homotypic immunity (for example PR8 primed, PR8 challenged) to which influenza-specific antibodies contribute (<xref ref-type="bibr" rid="bib47">Townsend et al., 1986</xref>; <xref ref-type="bibr" rid="bib9">Epstein and Price, 2010</xref>)<italic>.</italic> While the wild-type mice mounted a fast and strong recall response upon X31 challenge in PR8-primed mice (PR8 + X31), this was drastically diminished in T<italic>-Atg7</italic><sup><italic>−/−</italic></sup> mice in the lungs on day 5 (<xref ref-type="fig" rid="fig5">Figure 5A</xref>, absolute counts in <xref ref-type="fig" rid="fig5s1">Figure 5—figure supplement 1A</xref>). Even when three live virus immunizations were administered (PR8 primed, challenged with X31 then with PR8), T<italic>-Atg7</italic><sup><italic>−/−</italic></sup> mice were unable to mount recall responses. Surprisingly, T<italic>-Atg7</italic><sup><italic>−/−</italic></sup> mice survived the heterotypic viral challenges despite the vastly diminished numbers of secondary CD8<sup>+</sup> T<sub>eff</sub>.<fig-group><fig id="fig5" position="float"><object-id pub-id-type="doi">10.7554/eLife.03706.012</object-id><label>Figure 5.</label><caption><title><italic>Atg7</italic><sup>−/−</sup> memory CD8<sup>+</sup> T cells mount a significantly reduced recall response to secondary immunization.</title><p>(<bold>A</bold>) Recall responses to influenza. Here, WT and T-<italic>Atg7</italic><sup>−/−</sup> mice were split into three groups. In one, mice were immunized with 0.00032 HAU PR8 influenza and the NP-specific CD8<sup>+</sup> T cell response was measured in lungs on day 24 (PR8, n = 4). In another group, mice were immunized with 0.00032 HAU PR8 influenza followed by 0.32 HAU X31 influenza challenge on day 24. The recall response to this challenge was measured on day 5 post–challenge using NP-specific tetramers (PR8+X31, n = 6). In the third group, mice were immunized with 0.00032 HAU PR8 influenza, followed by 0.32 HAU X31 on day 24, and 30 days later immunized for a third time with 32 HAU PR8. The recall response to this third infection was measured in the lungs on day 5 post–challenge by tetramer (PR8 + X31 + PR8, n = 4). Quantification shows frequency of CD8<sup>+</sup> T cells that are NP-specific and are representative of two independent experiments. *p < 0.05, **p < 0.01, by Mann–Whitney U-test. (<bold>B</bold>) The <italic>Atg7</italic><sup><italic>−/−</italic></sup> CD8<sup>+</sup> T cell response to influenza vaccination. WT and T-<italic>Atg7</italic><sup>−/−</sup> mice were vaccinated twice, 22 days apart with 32 HAU of the live attenuated H1N1 vaccine, S-Flu. 30 days after the last vaccination, mice were challenged with 32 HAU X31 influenza. The CD8<sup>+</sup> T cell response to NP was measured in lungs on day 23 post–challenge by tetramer. As a control, unvaccinated mice were challenged with 32 HAU X31 and culled on day 4 due to weight loss and morbidity. Quantification indicates the percentage of CD8<sup>+</sup> T cells in the lung that are specific for NP on day 23 post–challenge. Example dot plots are shown. Data are representative of two independent experiments. *p < 0.05 by Mann–Whitney U-test (n = 4–5). (<bold>C</bold>) <italic>Atg7</italic><sup><italic>−/−</italic></sup> CD8<sup>+</sup> T cell kinetics to influenza vaccination. WT and T-<italic>Atg7</italic><sup>−/−</sup> mice were vaccinated with S-Flu twice, 22 days apart, and the CD8<sup>+</sup> T cell response to NP was tracked over time in blood by tetramer (n = 4–6). (<bold>D</bold>) <italic>Atg7</italic><sup><italic>−/−</italic></sup> CD8<sup>+</sup> T cell kinetics to influenza vaccination and challenge. WT and T-<italic>Atg7</italic><sup>−/−</sup> mice were vaccinated as described in (<bold>C</bold>). 30 days after the last vaccination, mice were challenged with 32 HAU X31 influenza. Using NP-specific tetramers, the CD8<sup>+</sup> T cell response to the vaccine regime and the challenge was tracked in the blood over time (n = 4–6). All values are mean ± s.e.m.</p><p><bold>DOI:</bold> <ext-link ext-link-type="doi" xlink:href="10.7554/eLife.03706.012">http://dx.doi.org/10.7554/eLife.03706.012</ext-link></p></caption><graphic xlink:href="elife03706f005"/></fig><fig id="fig5s1" position="float" specific-use="child-fig"><object-id pub-id-type="doi">10.7554/eLife.03706.013</object-id><label>Figure 5—figure supplement 1.</label><caption><title>Atg7<sup>−/−</sup> CD8<sup>+</sup> T cells fail to mount robust re-call responses to secondary infection.</title><p>(<bold>A</bold>) Recall CD8<sup>+</sup> T cell responses in WT and T-<italic>Atg7</italic><sup><italic>−/−</italic></sup> mice. Mice were immunized once with PR8 influenza (n = 4); twice with PR8+X31 influenza (n = 6); or three times with PR8 + X31 + PR8 (n = 4). Absolute counts of CD8<sup>+</sup> NP-tetramer<sup>+</sup> T cells were determined in the lungs at either day 24 (PR8), or day 5 post–challenge (PR8 + X31 & PR8 + X31 + PR8). All values are mean ± s.e.m.</p><p><bold>DOI:</bold> <ext-link ext-link-type="doi" xlink:href="10.7554/eLife.03706.013">http://dx.doi.org/10.7554/eLife.03706.013</ext-link></p></caption><graphic xlink:href="elife03706fs006"/></fig></fig-group></p><p>Using an established non-replicating pseudotyped influenza vaccination protocol (<xref ref-type="bibr" rid="bib35">Powell et al., 2012</xref>), we next tested whether H1N1 vaccinated T<italic>-Atg7</italic><sup><italic>−/−</italic></sup> mice could mount secondary CD8<sup>+</sup> T responses to heterotypic viral challenge. Mice were vaccinated twice, 22 days apart then challenged with X31 influenza. Large numbers of influenza NP-specific CD8<sup>+</sup> T cells were detected in the lungs of vaccinated wild-type mice 5 days post-challenge, but not in T<italic>-Atg7</italic><sup><italic>−/−</italic></sup> mice (<xref ref-type="fig" rid="fig5">Figure 5B</xref>). When the CD8<sup>+</sup> T cell kinetics to the vaccine regimen were followed over time in the blood of T<italic>-Atg7</italic><sup><italic>−/−</italic></sup> mice, influenza-specific CD8<sup>+</sup> T cells were detected at normal frequencies to primary immunization but the secondary ‘booster’ vaccine was unable to induce an increase in NP-specific CD8<sup>+</sup> T cell frequency like it could in wild-type mice (<xref ref-type="fig" rid="fig5">Figure 5C</xref>). Similarly, vaccinated T<italic>-Atg7</italic><sup><italic>−/−</italic></sup> mice failed to generate significant CD8<sup>+</sup> T cell responses to viral challenge in the blood (<xref ref-type="fig" rid="fig5">Figure 5D</xref>). As before, all mice survived the live viral challenge (data not shown). In summary, these data indicate that autophagy is required for the CD8<sup>+</sup> T cell recall response to either repeated immunizations with live virus or vaccination followed by live viral challenge. However, these results were not repeated in a mixed bone marrow chimera setting, making it difficult to exclude that lymphopenia, CD4<sup>+</sup> T cell help or CD4-expressing antigen-presenting cells might be responsible for defective recall response in T-<italic>Atg7</italic><sup><italic>−/−</italic></sup> mice.</p></sec><sec id="s2-6"><title>Rejuvenation of aged vaccine responses through autophagy</title><p>We reasoned that a natural setting where these findings might be applicable is the aged organism, where reduced autophagy levels are found in many organs and cell types (<xref ref-type="bibr" rid="bib43">Rubinsztein et al., 2011</xref>). Indeed, we have previously shown that autophagy levels are significantly reduced in the CD8<sup>+</sup> T cells of aged healthy human donors (>65 years) as compared to young donors (<30 years) (<xref ref-type="bibr" rid="bib34">Phadwal et al., 2012</xref>). Taking results presented here so far into account, we hypothesized that these low levels may contribute to the poor CD8<sup>+</sup> T<sub>mem</sub> formation and inefficient influenza vaccination observed in the elderly (<xref ref-type="bibr" rid="bib18">Kapasi et al., 2002</xref>; <xref ref-type="bibr" rid="bib12">Haynes et al., 2003</xref>; <xref ref-type="bibr" rid="bib19">Kedzierska et al., 2012</xref>).</p><p>We addressed whether diminished CD8<sup>+</sup> T<sub>mem</sub> formation can be improved in the elderly by modulating autophagy. First, we showed that mRNA levels of essential autophagy genes are decreased in sorted CD8<sup>+</sup> T cells from naïve old mice (2 years) as compared to young mice (8 weeks). The CD8<sup>+</sup> CD44<sup>hi</sup> memory compartment is particularly affected (<xref ref-type="fig" rid="fig6">Figure 6A</xref>). CD8<sup>+</sup> T cells from old mice also showed significantly decreased autophagic flux detected by counting LC3 spots in NP-specific CD8<sup>+</sup> T cells from young and old mice both in the presence and absence of an autophagy flux inhibitor (<xref ref-type="fig" rid="fig6">Figure 6B,C</xref>). This was confirmed by using two flow cytometry based autophagy detection, also in NP-specific CD8<sup>+</sup> T cells (<xref ref-type="fig" rid="fig6s1">Figure 6—figure supplement 1A and B</xref>).<fig-group><fig id="fig6" position="float"><object-id pub-id-type="doi">10.7554/eLife.03706.014</object-id><label>Figure 6.</label><caption><title>Boosting autophagy restores CD8<sup>+</sup> T cell responses to vaccination in elderly mice.</title><p>(<bold>A</bold>) Autophagy gene expression in CD8<sup>+</sup> T cells from young and elderly mice. CD44<sup>lo</sup> and CD44<sup>hi</sup> CD8<sup>+</sup> T cells were purified from 6 week old and 2 year old mice using fluorescent activated cell sorting. mRNA was extracted and the expression of essential autophagy genes was measured by q-PCR. Shown is the fold change in expression in CD8<sup>+</sup> T cells from old mice relative to expression in young mice (normalized to <italic>gapdh</italic> and <italic>hprt</italic>). (<bold>B</bold>) LC3 Spot count in CD8<sup>+</sup> T cells from young and old mice. Splenic CD8<sup>+</sup> T cells from 8 week old and 2 year old mice were treated with an autophagy flux inhibitor for 2 hr, as a control cells were left untreated. LC3 spot count was determined on CD8<sup>+</sup> CD44<sup>hi</sup> T cells using ImageStream. Representative images are shown (×60 magnification). **p = 0.0082, ***p = 0.0004 as determined by Student t-test (n = 4–5). (<bold>C</bold>) Quantification for images shown in (<bold>B</bold>). (<bold>D</bold>) Human T cell line Jurkat was incubated with 100 µM spermidine for 2, 4, or 6 hr or left untreated followed by whole protein extraction for LC3 Western Blot (upper panel). In the lower panel, Jurkat cells were incubated either with no spermidine (control), 100 µM, or 500 µM spermidine for 6 hr and then Western blotted for LC3. GAPDH was used as loading control for all Western blots. (<bold>E</bold>) Jurkat cells were treated with 1 mM DFMO or 1 mM DFMO with 1 µM spermidine for 48 hr or left untreated (control). In the final 6 hr of incubation, all cells were treated with 10 nM bafilomycin A1 and LC3-I to LC3-II conversion was assessed by Western Blot. (<bold>F</bold>) Jurkat cells were treated with 50 nM rapamycin, 100 µM or 500 µM spermidine for 6 hr followed by detection of phosphorylated S6 (Ser235/236) by Western Blot. As a control, cells were left untreated. (<bold>G</bold>) CD8<sup>+</sup> T cell kinetics to influenza vaccination in aged mice in the presence of spermidine. 8-week-old young WT and 23 month old WT and T-<italic>Atg7</italic><sup>−/−</sup> mice were vaccinated 22 days apart with S-Flu. 21 days prior to the first vaccination, aged WT and T-<italic>Atg7</italic><sup>−/−</sup> mice were administered spermidine in the drinking water at a concentration of 5 mM through to the experimental endpoint. As a control, 23 month old WT mice were administered water alone. The CD8<sup>+</sup> T cell response to NP was tracked over time in the blood by tetramer (n = 4–5). Y-axis depicts the frequency of CD8<sup>+</sup> T cells that are specific for NP. **p < 0.01 by Mann–Whitney U-test. (<bold>H</bold>) CD8<sup>+</sup> T cell kinetics to influenza vaccination and challenge in aged mice in the presence of spermidine. 8-week-old young WT and T-<italic>Atg7</italic><sup>−/−</sup> and 23 month old WT and T-<italic>Atg7</italic><sup>−/−</sup> mice were vaccinated as described in (<bold>G</bold>). 30 days after the last vaccination, mice were challenged with 32 HAU X31 and the CD8<sup>+</sup> T cell response to challenge was measured 9 days later in the lungs by tetramer. From 21 days prior to the first vaccination, through to the experimental end point, aged WT and T-<italic>Atg7</italic><sup>−/−</sup> mice were administered spermidine as before. Y-axis indicated the percentage of CD8<sup>+</sup> T cells that are specific for NP (n = 4–5). (<bold>I</bold>) CD8<sup>+</sup> T cell response to influenza challenge in vaccinated aged mice in the presence of spermidine. 9 days post–challenge, lungs were harvested and the NP-specific CD8<sup>+</sup> T cell response to influenza challenge was measured by tetramer. Example dot plots are gated on CD8<sup>+</sup> T cells. Bar chart shows absolute counts of NP-specific CD8<sup>+</sup> T cells in the lung per lobe. **p < 0.01, ****p < 0.0001 by Student <italic>t</italic> test (n = 4–5). All values are mean ± s.e.m.</p><p><bold>DOI:</bold> <ext-link ext-link-type="doi" xlink:href="10.7554/eLife.03706.014">http://dx.doi.org/10.7554/eLife.03706.014</ext-link></p></caption><graphic xlink:href="elife03706f006"/></fig><fig id="fig6s1" position="float" specific-use="child-fig"><object-id pub-id-type="doi">10.7554/eLife.03706.015</object-id><label>Figure 6—figure supplement 1.</label><caption><title>Autophagy levels are significantly diminished in antigen-specific CD8<sup>+</sup> T cells from aged mice.</title><p>Autophagy levels by CytoID (<bold>A</bold>) and LC3-II staining (<bold>B</bold>) in antigen-specific CD8<sup>+</sup> T cells from young and old mice. 8-week-old and 2 year old mice were immunized with PR8 influenza. On day 10 post-infection lungs were harvested and stained with CytoID or for LC3-II to assess autophagy levels in NP-Tetramer<sup>+</sup> CD8<sup>+</sup> T cells by flow cytometry. (<bold>A</bold>) ****p < 0.0001 (n = 7). (<bold>B</bold>) **p = 0.0056, (n = 7).</p><p><bold>DOI:</bold> <ext-link ext-link-type="doi" xlink:href="10.7554/eLife.03706.015">http://dx.doi.org/10.7554/eLife.03706.015</ext-link></p></caption><graphic xlink:href="elife03706fs007"/></fig></fig-group></p><p>Next, we chose the autophagy-inducing, naturally occurring compound spermidine to modulate autophagy in vivo, as it had been safely and effectively administered to mice previously (<xref ref-type="bibr" rid="bib8">Eisenberg et al., 2009</xref>). We first confirmed that spermidine induces autophagy in T cells in vitro in a dose and time-dependent manner shown by increased levels of LC3-II (<xref ref-type="fig" rid="fig6">Figure 6D</xref>). Spermidine levels in the blood are known to decrease with age (<xref ref-type="bibr" rid="bib38">Pucciarelli et al., 2012</xref>). To mimic this, we used the inhibitor of the natural spermidine synthesis pathway DFMO in the human Jurkat T cell line and found that autophagy levels detected by LC3 western blot significantly dropped in BafA treated samples and untreated samples (<xref ref-type="fig" rid="fig6">Figure 6E</xref>). Autophagy levels could be rescued by the addition of very low levels of spermidine in T cells (<xref ref-type="fig" rid="fig6">Figure 6E</xref>). As mTOR inhibition, such as achieved with rapamycin, is well known for its multiple side effects (<xref ref-type="bibr" rid="bib23">Lamming et al., 2013</xref>), we confirmed that spermidine induces autophagy in T cells in an mTOR-independent manner. Rapamycin completely abolished the phosphorylation of S6K, a major target downstream of activated mTOR, whereas spermidine had no such effect (<xref ref-type="fig" rid="fig6">Figure 6F</xref>). Finally, we administered spermidine to old and young mice via the drinking water at concentrations known to induce autophagy (<xref ref-type="bibr" rid="bib8">Eisenberg et al., 2009</xref>) during the influenza vaccination protocols as in <xref ref-type="fig" rid="fig5">Figure 5B–D</xref>. In response to first and second vaccination, and as observed in ageing humans, old mice do not mount a robust CD8<sup>+</sup> T cell response as compared to young mice. Spermidine dramatically enhanced influenza-specific CD8<sup>+</sup> T cell responses in old vaccinated wild-type mice but not in old vaccinated T<italic>-Atg7</italic><sup><italic>−/−</italic></sup> mice as tracked over time in blood (<xref ref-type="fig" rid="fig6">Figure 6G</xref>). Similarly, the response to live influenza challenge in the blood was improved twofold to threefold (back to levels similar to young mice) when spermidine was administered to old mice, but not in the absence of autophagy (<xref ref-type="fig" rid="fig6">Figure 6H</xref>). In the lungs, the number of NP-specific CD8<sup>+</sup> T cells responding to influenza challenge in vaccinated old mice were improved fivefold to sixfold in the presence of spermidine, but had little effect on the response of aged T<italic>-Atg7</italic><sup><italic>−/−</italic></sup> mice (<xref ref-type="fig" rid="fig6">Figure 6I</xref>).</p></sec></sec><sec id="s3" sec-type="discussion"><title>Discussion</title><p>Hematopoietic and immune health both decline significantly with age (<xref ref-type="bibr" rid="bib4">Beerman et al., 2010</xref>). With the ageing population, this has become a substantial health and socio-economic problem for the developed world. Our work sheds light on the cellular mechanisms of immune ageing and how to improve responses to vaccination in the elderly.</p><p>The naïve T cell compartment in the absence of several different essential autophagy genes has been extensively analysed (<xref ref-type="bibr" rid="bib39">Puleston and Simon, 2014</xref>). These models uniformly demonstrated increased damaged mitochondria leading to cell death and peripheral lymphopenia. In this study, we show how this lymphopenia drives the remaining T cells to take on an activated/memory phenotype as they proliferate in an attempt to re-fill the T cell niche, often termed ‘virtual memory’ (<xref ref-type="bibr" rid="bib16">Jia and He, 2011</xref>). Thus, we provide a link between lymphopenia and the formation of the ‘virtual memory’ compartment and highlight the significant changes in T cell surface phenotype that can occur in a lymphopenic environment. Interestingly, the virtual memory compartment is also a hallmark of the ageing adaptive immune system. This observed phenotype in naïve mice would have precluded a coherent analysis of antigen-activated T cells; however, the use of mixed BM chimeras crucially excluded a contribution of the skewed naïve repertoire to the CD8<sup>+</sup> T<sub>mem</sub> phenotype.</p><p>In this study, we showed a novel and essential link for autophagy in the formation of CD8<sup>+</sup> T<sub>mem</sub> to infection. The absence of autophagy also dramatically altered the ability of CD8<sup>+</sup> T cells to respond to secondary infection. Interestingly, the severely depleted CD8<sup>+</sup> T cell recall response did not prevent T-<italic>Atg7</italic><sup>−/−</sup> mice from surviving a lethal secondary heterotypic influenza challenge. Influenza virus is typically controlled by both antibodies and cellular immunity. However, in a heterotypic challenge, the surface hemagglutinin and neuraminidase (recognized by antibodies) differ between the two heterotypic strains, while the viral internal proteins (presented by MHC class I to CD8<sup>+</sup> T cells) remain the same, leaving the CD8<sup>+</sup> T cells to provide protection (<xref ref-type="bibr" rid="bib41">Rimmelzwaan et al., 2007</xref>). The unique protective role of CD8<sup>+</sup> T cells in heterotypic immunity was described several decades ago in mice (<xref ref-type="bibr" rid="bib41">Rimmelzwaan et al., 2007</xref>). The observation that the majority of CD8<sup>+</sup> T cell epitopes are cross-reactive between subtypes and are located in the most conserved regions of the internal proteins (nucleoprotein and matrix protein) confirms their role. In humans, the evidence, typical for human studies, is sparse and more circumstantial. A study from 1983 by McMichael et al demonstrated that in experimentally infected individuals, virus-specific cytotoxicity inversely correlated with the extent of virus shedding in the absence of virus-specific antibodies for the strain that was used for infection (<xref ref-type="bibr" rid="bib27">McMichael et al., 1983</xref>). More recently, Lalvani et al found higher frequencies of pre-existing T cells to conserved epitopes in individuals who developed less severe illness in the 2009 H1N1 flu pandemic in the absence of cross-protective antibodies (<xref ref-type="bibr" rid="bib45">Sridhar et al., 2013</xref>). We would postulate therefore that the survival of the T-<italic>Atg7</italic><sup>−/−</sup> mice after the secondary heterotypic challenge relies on the very few effector CD8<sup>+</sup> T cells generated. These data also suggest that eliciting CD8<sup>+</sup> T cell memory responses in influenza vaccination is highly desirable, particularly to protect from pandemic flu.</p><p>This also raises interesting questions as to why such robust secondary T cell responses have evolved when survival can be achieved with a fraction of what is observed normally in nature. Although no CD8<sup>+</sup> T<sub>mem</sub> cells can be detected by conventional tetramer technology 20–30 days post-immunization in T-<italic>Atg7</italic><sup>−/−</sup> mice, some memory cells must remain below the detection limit at very low frequency. In addition, these <italic>Atg7</italic><sup>−/−</sup> CD8<sup>+</sup> T cells must maintain normal functional and proliferative capacity as they are detected at early time points after secondary immunization, characteristic of true memory T cell kinetics.</p><p>To explain how autophagy affects the maintenance of CD8<sup>+</sup> T<sub>mem</sub> cells, we addressed two alternative hypotheses experimentally: (1) accumulation of cellular damage affects the long-lived CD8<sup>+</sup> T<sub>mem</sub> in particular and (2) autophagy is required for the switch from glycolysis to mitochondrial respiration important for the survival of CD8<sup>+</sup> T<sub>mem</sub> cells. Regarding the first one, we show evidence for the accumulation of mitochondria and ROS in the absence of autophagy. However, a limitation of our study is, we did not test for accumulation of other organelles and protein aggregates. As for the second hypothesis, initial studies involved TRAF6 (TNFR-associated factor 6) in the regulation of CD8<sup>+</sup> T<sub>mem</sub> development by modulating fatty acid metabolism and thereby increasing mitochondrial respiration (<xref ref-type="bibr" rid="bib33">Pearce et al., 2009</xref>; <xref ref-type="bibr" rid="bib48">van der Windt et al., 2012</xref>). Interestingly, TRAF6, as an E3 ubiquitin ligase, has recently been shown to stabilize essential proteins in the autophagy pathway such as Ulk1 (<xref ref-type="bibr" rid="bib32">Nazio et al., 2013</xref>) and Beclin-1 (<xref ref-type="bibr" rid="bib44">Shi and Kehrl, 2010</xref>). We therefore hypothesize that TRAF6 controls metabolism via autophagy and propose three mechanisms contributing to autophagy's role in mitochondrial respiration: (1) through lipophagy, the degradation of lipids (<xref ref-type="bibr" rid="bib50">Weidberg et al., 2009</xref>), autophagy provides the fatty acids that fuel the Krebs cycle with Acetyl-CoA though β-oxidation, (2) through degradation of glycolytic enzymes (<xref ref-type="bibr" rid="bib51">Xiong et al., 2011</xref>), and (3) through mitophagy, autophagy provides mitochondrial quality control that is required for functional mitochondrial ATP generation. We provide evidence for the latter mechanism here. This suggests that autophagy is important in the control of metabolic pathways in T cells. Interestingly both the accumulation of damaged mitochondria/oxidative stress as well as a bias to preferentially use glycolysis are hallmarks of the ageing cell (<xref ref-type="bibr" rid="bib13">Hipkiss, 2006</xref>).</p><p>In this study, we showed that autophagy levels are impaired in CD8<sup>+</sup> T cells from aged mice. Using spermidine, we could dramatically improve the CD8<sup>+</sup> T cell response to vaccination and infection in elderly mice in an autophagy-dependent manner. However, aged CD4<sup>+</sup> T cells also undergo immune senescence. While inducing autophagy with spermidine in the CD8<sup>+</sup> T cell compartment clearly contributes to the increased numbers of antigen-specific CD8<sup>+</sup> T cells, the limitation of these experiments is that we cannot gauge the contribution of improved CD4<sup>+</sup> T cell function to CD8<sup>+</sup> T cell numbers. This will be included in future studies.</p><p>These results offer the first cell-intrinsic explanation as to why T cell memory is defective in old age. Spermidine is a pleiotropic compound and its major function has been described in yeast as an inhibitor of histone acetyl transferases, inducing epigenetic changes to autophagy-related gene expression (<xref ref-type="bibr" rid="bib8">Eisenberg et al., 2009</xref>). We showed that spermidine operates independently of mTOR to induce autophagy. These findings offer the prospect of improving vaccine T cell responses in the elderly through spermidine in an mTOR-independent fashion. However, no other ways of inducing autophagy were tested here. We expect that fasting or other mTOR-independent drugs such as resveratrol (<xref ref-type="bibr" rid="bib29">Morselli et al., 2009</xref>; <xref ref-type="bibr" rid="bib30">Morselli et al., 2010</xref>) may also be beneficial for CD8<sup>+</sup> T<sub>mem</sub> maintenance. However, spermidine is significantly more attractive than previously published T<sub>mem</sub>-boosting compounds such as metformin (<xref ref-type="bibr" rid="bib33">Pearce et al., 2009</xref>) and rapamycin (<xref ref-type="bibr" rid="bib3">Araki et al., 2009</xref>), that come with unwelcome side effects and toxicity in humans. The elucidation of the precise role of spermidine will be the subject of future work and will enable its development as a safe, readily administrable immune-modulating drug.</p></sec><sec id="s4" sec-type="materials|methods"><title>Material and methods</title><sec id="s4-1"><title>Mice</title><p><italic>CD4-Cre</italic> mice (<xref ref-type="bibr" rid="bib24">Lee et al., 2001</xref>) were from Adeline Hajjar (University of Washington) and crossed with <italic>Atg7</italic><sup><italic>flox/flox</italic></sup> (<xref ref-type="bibr" rid="bib22">Komatsu et al., 2005</xref>) to obtain T-<italic>Atg7</italic><sup>−/−</sup> mice (<italic>CD4-Cre</italic><sup><italic>+</italic></sup> <italic>Atg7</italic><sup><italic>+/+</italic></sup>) on a C57BL/6 background. Unless otherwise stated, all mice were 6–12 weeks of age at the start of each experiment and were age and sex matched. <italic>CD4-Cre</italic><sup><italic>−</italic></sup> <italic>Atg7</italic><sup><italic>+/+</italic></sup> mice were used as wild-type controls and were littermates where possible. No phenotype was observed in <italic>CD4-Cre</italic><sup><italic>+</italic></sup> <italic>Atg7</italic><sup><italic>+/−</italic></sup> mice. Old mice and young control mice were purchased from Charles River, UK. C57BL/6 SJL CD45.1 mice for bone marrow chimeras were purchased from Biomedical Services, Oxford. All mice were housed in Biomedical Services, Oxford and animal experiments were approved by the local ethical review committee and performed under UK project license (PPL 39/2809).</p></sec><sec id="s4-2"><title>Flow cytometry</title><p>The following antibodies were used for flow cytometry (antibody clone in brackets): CD8 (Ly2) PE/PE-CY7; CD8 (53-6.7) FITC/eF450/PE-Cy7; CD4 (GK1.5) PE/FITC/APC; TCRβ (H57-597) FITC/PE/PE-Cy7/APC; CD3 (145-2C11) eF450/APC; CD62L (MEL-14) FITC/PE-Cy7; CD44 (IM7) FITC/PE-Cy7; KLRG1 (2F1) FITC/PE-Cy7; IRF4 (3E4) FITC; EOMES (Dan11mag) PE-Cy7; CD127 (A7R34) FITC/PE/eF450; Ki-67 (SolA15) eF450; CD24 (M1/69) APC; CD16/32 (93) purified Fc block; CD45.2 (104) eF450; all from eBioscience (San Diego, CA, USA). PD-1 (29F.1A12) PE-Cy7; PD-1 (RMP1-30) PE; CD45.2 (104) PE-Cy7; TIM-3 (B8.2C12) PE/APC; Bcl-2 (BCL/10C4) PE; all from Biolegend (San Diego, CA, USA). Glut1 (q) PerCP; IL-15Rα (BAF551) biotinylated; both from R&D Systems (Minneapolis, USA).</p><p>For surface staining, cells were suspended in PBS, 2% FCS, 5 mM EDTA and stained at 4°C. Anti-CD16/32 (Fc Block, eBioscience) was generally added to antibody mix to minimize non-specific staining. LIVE/DEAD Fixable Violet Dead Cell Stain Kit (Life Technologies) was used prior to surface staining to exclude dead cells. For cytoplasmic intracellular staining (Bcl-2), cells were stained for surface antibodies then fixed in IC fixation buffer before permeabilisation with perm buffer (eBioscience). Cells were then suspended in perm buffer at RT for antibody staining. For nuclear targets (IRF, EOMES, Ki-67), after surface staining cells were fixed and permeabilised with FoxP3 Fixation/Permeabilisation kit (eBioscience) before resuspension in perm buffer for intracellular staining at RT.</p><p>Absolute cell counts were performed on peripheral blood taken from the lateral tail vein of live animals, collected in heparin-coated tubes (Microvette 300, Sarstedt [Nümbrecht, Germany]) to avoid coagulation. Cell counts were calculated with BD TruCount tubes (BD Bioscience, NJ, USA) according to the manufacturer's instructions.</p><p>Tetramers were generated as previously described (<xref ref-type="bibr" rid="bib2">Altman et al., 1996</xref>) and MHC-Class I monomers stored at −80°C. Biotinylated monomers were tetramerized with Streptavidin PE or APC at the right concentration to achieve 1:1 ratio with biotin binding sites and added in 1/10th volumes waiting 10 min between additions. Tetramerized complexes were stored at 4°C. Peptide sequences for MCMV tetramers were as follows: m45 <sup>985</sup>HGIRNASFI<sup>993</sup>, H-2D<sup>b</sup>-restricted; m38 <sup>316</sup>SSPPMFRV<sup>323</sup>, H-2K<sup>b</sup>-restricted; IE3 <sup>416</sup>RALEYKNL<sup>423</sup>, H-2K<sup>b</sup>-restricted. The peptide sequence for the influenza tetramer was as follows: NP <sup>366</sup>ASNENMETM<sup>374</sup>, H-2D<sup>b</sup>-restricted. Cells were always stained with tetramers prior to surface antibody staining in PBS, 2% FCS at 37°C for 15 min. Tracking tetramer-positive cells over time in the blood was performed by serially bleeding the lateral tail vein. After erythrocyte lysis with red cell lysis buffer (eBioscience), remaining white cells were stained with tetramer and surface antibodies as described above.</p><p>Autophagy detection by flow cytometry was measured using the CytoID Autophagy Detection Kit (Enzo Life Science, Exeter, UK). Splenocytes and cells isolated from the lung were first stained with CytoID according to the manufacturer's instructions, prior to tetramer and surface antibody staining. Autophagy was also measured using flow cytometry by quantifying LC3-II mean fluorescence intensity using the FlowCellect Autophagy LC3 Antibody-based Assay Kit (FCCH100171, Merk-Millipore, MA, USA) according to the manufacture's instructions and always following cell surface antibody staining. Use of this kit includes a step where cytosolic LC3-I is washed from the cell, leaving only membrane bound LC3-II prior to staining.</p><p>For apoptosis detection, splenocytes were stained with m45-tetramer and surface antibody as described above and then stained with LIVE/DEAD Fixable Violet Dead Cell Stain Kit. Cells were finally stained with Annexin V PE-Cy7 (eBioscience) in Annexin V binding buffer at room temperature. Apoptotic cells were determined as LIVE/DEAD cell dye negative (live cells), Annexin V positive.</p><p>GLUT-1 was measured through binding to its ligand, the receptor binding domain (RBD) of a recombinant glycoprotein from the human T lymphotrophic virus (HTLV) fused to eGFP (H<sub>RBD</sub>-eGFP) (<xref ref-type="bibr" rid="bib28">Montel-Hagen et al., 2008</xref>).</p><p>For mitochondrial mass analysis, cells were stained with MitoTracker Green (Life Technologies, Carlsbad, CA, USA) at 150 nM in PBS, 2% FCS for 30 min at 37°C after tetramer and surface antibody staining. To measure mitochondrial superoxide production, MitoSOX Red (Life Technologies) was used at final concentration of 5 µM from a 5 mM stock in PBS, 2% FCS for 30 min at 37°C following tetramer and cell surface staining. All flow cytometry experiments were performed on a Cyan flow cytometer (Beckman Coulter, Brea, CA, USA).</p></sec><sec id="s4-3"><title>Bone marrow (BM) chimera</title><p>BM cells were extracted from a single 8-week old wild-type, T-<italic>Atg7</italic><sup><italic>−/−</italic></sup> (both CD45.2<sup>+</sup>), and C57BL/6 SJL mouse expressing CD45.1. After erythrocyte lysis, 3 × 10<sup>6</sup> wild-type or T-<italic>Atg7</italic><sup><italic>−/−</italic></sup> BM cells were added to 3 × 10<sup>6</sup> CD45.1+ BM cells (1:1 CD45.2<sup>+</sup>:CD45.1<sup>+</sup>) in a total volume of 200 µl PBS. The 1:1 BM mix was injected i.v 2 hr after lethal irradiation (450 cGy twice, 4 hr apart) to C57BL/6 SJL CD45.1+ recipients (total of 6 × 10<sup>6</sup> BM cells per mouse in 200 µl). After 8 weeks of reconstitution, mixed BM chimera was immunized with MCMV or PR8 influenza, or left unimmunized. WT and T-<italic>Atg7</italic><sup><italic>−/−</italic></sup> mice were always used as controls.</p></sec><sec id="s4-4"><title>Immunizations</title><p>Mice were immunized i.v. with 1 × 10<sup>6</sup> p.f.u MCMV in 100 µl (Smith strain ATCC: VR194). Unimmunized controls were injected with 100 µl PBS i.v. For influenza infections, mice were administered intra-nasally with either A/PR/8/34 (PR8, H1N1 Cambridge) influenza or X31 (H3N2) influenza at the stated dose in 50 µl viral dilution media (VDM; DMEM, 0.1% BSA, 10 mM HEPES, 100 U/ml penicillin, 100 µg/ml streptomycin, and 2 mM glutamine; all from Sigma, St Louis, MO, USA). Mice were anaesthetized with isofluorane and droplets of VDM-containing virus were applied to the nares until the total 50 µl was inhaled. Unimmunized mice were always used as controls (50 µl of VDM alone). For influenza vaccination, mice were immunized intra-nasally with 32 HAU pseudotyped H1N1 influenza (S-Flu) (<xref ref-type="bibr" rid="bib35">Powell et al., 2012</xref>), using the influenza infection protocol described above, and challenged with 32 HAU X31. Unvaccinated and fully naïve mice were used as controls in all influenza vaccine experiments. In unvaccinated mice that received X31 challenge, a combination of weight loss and clinical score was used as a humane endpoint.</p></sec><sec id="s4-5"><title>q-PCR</title><p>Cells were purified with a MoFlo cell sorter (Beckman Coulter) by their surface markers. RNA was extracted using RNeasy Kit (Qiagen, Hilden, Germany) and quantified using a Nanodrop spectrophotometer (Thermo Scientific, Waltham, MA, USA). RNA was reverse transcribed (RT) using a High Capacity RNA to cDNA kit (Applied Biosystems (AB), Foster City, CA, USA). Resulting cDNA was stored at −20°C. Real-time quantitative PCR using comparative Ct method (ΔΔCt) was utilized to evaluate gene expression using validated TaqMan probes (AB) on a 7500 Fast Real-time PCR machine (AB). Conditions: (1) 50°C, 2 min; (2) 95°C, 10 min; (3) 95°C 15 s; (4) 60°C 1 min; 40 cycles of 3–4. The assay IDs for the primers of the analyzed genes are as follows: Mm00504340_m1 (<italic>Atg5</italic>), Mm00512209_m1 (<italic>Atg7</italic>), Mm01264428_m1 (<italic>Atg9</italic>), Mm00470550_m1 (<italic>Atg10</italic>), Mm00503201_m1 (<italic>Atg12</italic>), Mm0051717_m1 (<italic>beclin1</italic>), Mm00458724_m1 (<italic>Map1lc3a</italic>), Mm01545399_m1 (<italic>hprt</italic>), Mm99999915_g1 (<italic>gapdh</italic>).</p></sec><sec id="s4-6"><title>ImageStream</title><p>ImageStream (Amnis imaging flow cytometer, MA, USA) has previously been used to determine autophagic flux (<xref ref-type="bibr" rid="bib34">Phadwal et al., 2012</xref>). To determine LC3 spot count, we stained cells for LC3-II (following cell surface staining) using the FlowCellect Autophagy LC3 Antibody-based Assay Kit (FCCH100171, Merk-Millipore) according to the manufacture's instructions. Before LC3 detection, to assess autophagic flux, cells were incubated for 2 hr with the autophagy flux inhibitor provided by the kit specified and according to the manufacturer's instructions. LC3 spot count was quantified using Ideas software (Amnis), which contains a specialised objective spot counting feature. Images collected were all at ×60 magnification.</p></sec><sec id="s4-7"><title>Western blot</title><p>3 × 10<sup>6</sup> Jurkat cells (human CD4<sup>+</sup> T cell line) or 5 × 10<sup>6</sup> purified primary CD8<sup>+</sup> T cells were lysed on ice using 100 μl 1 × NP-40 Lysis Buffer. Protein concentration in supernatant was measured using BCA Protein Assay Kit (PI-23227, Thermo Scientific), and reducing Laemmli Sample Buffer was added to make protein samples for SDS-PAGE. Proteins of 30–50 μg per lane were separated on 16% SDS-PAGE and transferred to PVDF membrane (IPFL00010, Millipore, MA, USA). After blocking with 5% skim milk, the membrane was blotted using the following primary antibodies: LC3 (L8918, Sigma) (1:1000), GAPDH (MAB374, Millipore) (1:10,000), pS6 (2211, Cell Signaling Technology, Danvers, MA, USA) (1:5000). For S6 blotting, membranes were blotted with anti-pS6 first, then stripped for re-blot using primary antibody against S6 (2217, Cell Signaling Technologies) (1:1000). IRDye secondary antibodies were bought from LI-COR (Lincoln, NE, USA): (926-32,211, 926-68,020) (1:15,000). For the purification of primary CD8<sup>+</sup> T cells, cells were negatively selected from splenocytes using the CD8<sup>+</sup> T cell Isolation kit (130-104-075, Miltenyi, Bergisch Gladbach, Germany) according to the manufacturer's instructions.</p></sec><sec id="s4-8"><title>Influenza viral titers (TCID<sub>50</sub> and hemagglutination)</title><p>Determination of the 50% tissue culture infective dose (TCID<sub>50</sub>) was performed on MDCK-SIAT1 cells in a 96-well flat-bottom plate performed by serial dilution of lung homogenates onto 3 × 10<sup>4</sup> MDCK-SIAT1 cells followed by incubation for 72 hr at 37°C. Virus was then detected by hemagglutination, where 50 µl 1% (vol/vol) human erythrocytes (adjusted so that a 1:2 dilution gave an optical density at 600 nm) was added to 50 µl of the serially diluted lung homogenates/MDCK-SIAT1 supernatant in a 96-well V-bottom plate. Hemagglutination was analyzed by the loss of teardrop formation after tilting the plate. TCID<sub>50</sub> was calculated as described by <xref ref-type="bibr" rid="bib40">Reed and Muench (1938)</xref>.</p></sec><sec id="s4-9"><title>Statistical analyses</title><p>All data were presented as mean ± s.e.m. p values were determined using either a Mann–Whitney U-test or two-tailed Student's <italic>t</italic> test using GraphPad Prism software. Significant statistical differences are indicated in the figure legends or on the graphs themselves.</p></sec><sec id="s4-10"><title>In vivo spermidine treatment</title><p>Mice were administered spermidine in the drinking water 21 days prior to immunization through to the experimental endpoint. Spermidine was added at a concentration of 5 mM from a 1 M aqueous stock. Spermidine-containing drinking water was replenished every 2–3 days.</p></sec></sec></body><back><ack id="ack"><title>Acknowledgements</title><p>The study was funded by the MRC Human Immunology Unit and the Wellcome Trust. The Chinese Scholarship Council is gratefully acknowledged for the stipend and fees of HZ; the Lady Tata fund for AW's stipend and fees; and TJP is funded by the Townsend-Jeantet Trust (registered charity no. 1011770) and the MRC Human Immunology Unit. Thanks also goes to Rosanna McEwen-Smith for assistance in setting up the influenza studies.</p></ack><sec sec-type="additional-information"><title>Additional information</title><fn-group content-type="competing-interest"><title>Competing interests</title><fn fn-type="conflict" id="conf1"><p>The authors declare that no competing interests exist.</p></fn></fn-group><fn-group content-type="author-contribution"><title>Author contributions</title><fn fn-type="con" id="con1"><p>DJP, Conception and design, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article</p></fn><fn fn-type="con" id="con2"><p>HZ, Acquisition of data, Analysis and interpretation of data</p></fn><fn fn-type="con" id="con3"><p>EL, Acquisition of data, Analysis and interpretation of data</p></fn><fn fn-type="con" id="con4"><p>IP, Acquisition of data, Analysis and interpretation of data</p></fn><fn fn-type="con" id="con5"><p>ASW, Acquisition of data, Analysis and interpretation of data</p></fn><fn fn-type="con" id="con6"><p>TJP, Conception and design, Acquisition of data, Analysis and interpretation of data, Contributed unpublished essential data or reagents</p></fn><fn fn-type="con" id="con7"><p>SS, Acquisition of data, Contributed unpublished essential data or reagents</p></fn><fn fn-type="con" id="con8"><p>VC, Analysis and interpretation of data, Contributed unpublished essential data or reagents</p></fn><fn fn-type="con" id="con9"><p>ARMT, Conception and design, Analysis and interpretation of data, Contributed unpublished essential data or reagents</p></fn><fn fn-type="con" id="con10"><p>PK, Conception and design, Analysis and interpretation of data, Contributed unpublished essential data or reagents</p></fn><fn fn-type="con" id="con11"><p>AKS, Conception and design, Analysis and interpretation of data, Drafting or revising the article</p></fn></fn-group><fn-group content-type="ethics-information"><title>Ethics</title><fn fn-type="other"><p>Animal 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pub-id-type="doi">10.1038/267354a0</pub-id></element-citation></ref></ref-list></back><sub-article article-type="article-commentary" id="SA1"><front-stub><article-id pub-id-type="doi">10.7554/eLife.03706.016</article-id><title-group><article-title>Decision letter</article-title></title-group><contrib-group content-type="section"><contrib contrib-type="editor"><name><surname>Levine</surname><given-names>Beth</given-names></name><role>Reviewing editor</role><aff><institution>UT Southwestern Medical School</institution>, <country>United States</country></aff></contrib></contrib-group></front-stub><body><boxed-text><p>eLife posts the editorial decision letter and author response on a selection of the published articles (subject to the approval of the authors). An edited version of the letter sent to the authors after peer review is shown, indicating the substantive concerns or comments; minor concerns are not usually shown. Reviewers have the opportunity to discuss the decision before the letter is sent (see <ext-link ext-link-type="uri" xlink:href="http://elifesciences.org/review-process">review process</ext-link>). Similarly, the author response typically shows only responses to the major concerns raised by the reviewers.</p></boxed-text><p>Thank you for sending your work entitled “Autophagy as a critical regulator of CD8+ T cell memory and a route to improved vaccination in the elderly” for consideration at <italic>eLife.</italic> Your article has been favorably evaluated by Tadatsugu Taniguchi (Senior editor), a member of the Board of Reviewing Editors, and 2 expert reviewers in your field.</p><p>The Reviewing editor and the reviewers discussed their comments before we reached this decision, and the Reviewing editor has assembled the following comments to help you prepare a revised submission.</p><p>The reviewers and Reviewing editor all feel that, overall, this is an interesting, well-performed study addressing the role of autophagy and ageing effects on autophagy in viral-specific CD8 memory T cells. We think that this study has important potential implications for understanding the role of autophagy in a crucial basic aspect of immunology, T cell memory, and, as pointed out by the authors, has potential clinical implications for improving vaccine responses in the elderly.</p><p>The reviewers have made several suggestions to further improve the manuscript. We are enclosing a list of the major points. Some of the points request further text explanations or modifications of the data analysis. Other points suggest a need for some additional data; however, as a general principle, given the overall enthusiasm for the work and the editorial policies of <italic>eLife</italic>, if these points cannot be quickly addressed by new experimental data, we ask that you revise the text (Discussion) to acknowledge the limitations of the present study.</p><p>One exception to this principle is point #8; although we recognize that your lab has previously published studies using the BDS feature to measure autophagic flux in lymphocytes, we do believe that, prior to publication, this assay should be confirmed by at least one additional autophagic flux assay to more convincingly demonstrate a defect in autophagic flux in T/Atg7-/- cells. Another exception is point #9; the LC3 assays in <xref ref-type="fig" rid="fig6">Figure 6D and 6E</xref> are performed in the same cells; the difference in the blots for control cells suggests a technical problem that should be resolved prior to publication.</p><p>1) The authors have not directly examined the effects of memory CD8 T cells on viral clearance and/or protection against pathological lung damage during influenza virus. If the authors have data in the adoptive transfer experiments in <xref ref-type="fig" rid="fig3">Figure 3</xref> evaluate the functions of wild-type and Atg7-/- memory T on viral clearance and/or lung damage, such data should be included in the manuscript. If they have not collected such data in their experiments, they should acknowledge this limitation.</p><p>2) The authors use a model of heterotypic immunity against influenza to avoid a model in which antibody-mediated responses are involved in protection against viral challenge. However, it is notable that mouse survival is not affected by autophagy deficiency in T cells. The reviewers ask whether the authors have tried to use higher titers of the viruses or lethal strains such as those described before (Virology 2011, 412:36-45). If antibodies are the main effectors against influenza and CD8 T cells are not sufficient or required for the protection against influenza challenge, this should be clarified by the authors. In addition, the authors should more fully discuss the data on the role of human CD8 T cell responses in protection against influenza challenge after vaccines. The argument that the findings in mice may have relevance to diminished vaccine responses to flu vaccines in elderly relies on the relevance of CD8 T cell memory responses to vaccine efficacy. The final paragraph mentions briefly that CD8 T cell responses are an important correlate of vaccine protection; however, this should be discussed more extensively in light of the lack of in-depth experimental evidence in this study in the mouse for a role of memory CD8 T cells in protection against disease caused by viral challenge.</p><p>3) T/Atg7-/- mice show defects in T cell development and a reduction of CD4 and CD8 T cells in the spleen and lymph nodes (J. Immunol. 182:4046-4055 and <xref ref-type="fig" rid="fig2">Figure 2</xref>). In <xref ref-type="fig" rid="fig1">Figure 1</xref>, the authors should show the percentage and total numbers of antigen-specific CD8 T cells among all cells in the lung, not just among gated CD8 T cells. The authors conclude that there is “normal expansion of the antigen-specific effector CD8+ T cell compartment”. However, a general lymphopenia in CD4 and CD8 T cells in these mice should be clearly presented in <xref ref-type="fig" rid="fig1">Figure 1</xref>, although a comparable fold of CD8 expansion was observed.</p><p>4) The reviewers feel that the interpretation of data in <xref ref-type="fig" rid="fig5">Figure 5</xref> is complicated by lymphopenia in T/Atg7-/- mice as well as autophagy deficiency in the CD4 T cell compartment. This point needs to be addressed in the authors' response.</p><p>5) In <xref ref-type="fig" rid="fig6">Figure 6</xref>, the authors should consider immunodeficiency in aged CD4 T cells (Wakikawa et al., Exp Gerontol 34, 231-242; Cheung et al., Exp Gerontol 18, 451-460). Spermidine would target all cells types in vivo, including CD4+ T helper cells. The experiments performed do not distinguish between these possibilities.</p><p>6) The knockout of essential genes from the autophagic cascade entails a major cellular dysfunction due to accumulating, dysfunctional mitochondria (and perhaps other organelles and protein aggregates). Hence, the phenotype of Atg7 knockout cells is not just the result of autophagy inhibition but rather that of accumulating damage. This problem should be discussed. It would be even better to include some of the data that are mentioned as “not shown” with regard to the phenotype of mice lacking Atg5 in T cells.</p><p>7) The authors only use one autophagy inducer, spermidine, in their experiments. Hence, it is still possible that autophagy would be only one of the spermidine effects (knowing that spermidine that might mediate additional metabolic effects on polyamine metabolism) that accounts for the improvement of CD8 memory T cells. The paper would be more complete, if the authors used other autophagy inducers (such as resveratrol, fasting etc). At a minimum, the current limitation should be discussed.</p><p>8) Autophagic flux is only measured by the 'BDS feature', namely the co-localization of a lysosomal marker and LC3, measured by means of the Amnis Imagestream system. It is important to confirm these measurements (and in particular the efficiency of the ATG7 knockout) by immunoblot detection of the LC3 lipidation in the absence and presence of autophagy inhibitors.</p><p>9) The controls in <xref ref-type="fig" rid="fig6">Figure 6d and 6e</xref> look very different with regard to the proportion of LC3-I and LC3-II. This could be due to several different factors, including differences in basal autophagy conditions (density, nutritional factors) or sample processing (i.e. freeze-thawing samples will result in degradation of LC3-I which is more labile). The discrepancy in experimental conditions that results in such a notable discrepancy in the proportion of LC3-I and LC3-II in controls of the same cell type in different figures should be resolved.</p></body></sub-article><sub-article article-type="reply" id="SA2"><front-stub><article-id pub-id-type="doi">10.7554/eLife.03706.017</article-id><title-group><article-title>Author response</article-title></title-group></front-stub><body><p><italic>1) The authors have not directly examined the effects of memory CD8 T cells on viral clearance and/or protection against pathological lung damage during influenza virus. If the authors have data in the adoptive transfer experiments in</italic> <xref ref-type="fig" rid="fig3"><italic>Figure 3</italic></xref> <italic>evaluate the functions of wild-type and Atg7-/- memory T on viral clearance and/or lung damage, such data should be included in the manuscript. If they have not collected such data in their experiments, they should acknowledge this limitation</italic>.</p><p>We have measured viral titers against influenza in the lungs of normal T-Atg7-/- mice (but not bone marrow chimeras as in <xref ref-type="fig" rid="fig3">Figure 3</xref>). While viral titers in the lungs of T-Atg7-/-mice are comparable to WT mice at day 3 of infection, they are significantly higher on day 6, suggesting T-Atg7-/- mice clear virus less efficiently than WT mice. This is probably due to the significantly lower absolute count of effector antigen-specific CD8 T cells. We expect that T-Atg7-/- mice would eventually clear the influenza virus as they survive the influenza challenge. The reason we did not measure this in a bone marrow chimera or adoptive transfer setting is because both the Atg7+/+ CD45.1 and Atg7-/- CDD45.2+ CD8 T cells would contribute toward viral clearance and it would be impossible to discern either's contribution. However, an additional mouse model as suggested by the reviewer, i.e. adoptive transfer experiments of Atg7-/- T cells into a T cell depleted host, would probably address this issue. This experiment could not be accomplished in the time frame given. We have now added the graph of viral clearance as <xref ref-type="fig" rid="fig1">Figure 1g</xref> to the manuscript.</p><p><italic>2) The authors use a model of heterotypic immunity against influenza to avoid a model in which antibody-mediated responses are involved in protection against viral challenge. However, it is notable that mouse survival is not affected by autophagy deficiency in T cells. The reviewers ask whether the authors have tried to use higher titers of the viruses or lethal strains such as those described before (Virology 2011, 412:36-45)</italic>.</p><p>We too have thought about this experiment. However, even in wild type mice it is possible to overcome heterotypic immunity with a suitably intense secondary challenge. Furthermore, the dose we used is already at least 10,000 morbid doses and therefore, we think using a different strain also at a highly lethal dose is unlikely to give different results. Because of this, we feel that doing this experiment in knock out animals would not add any information on how these animals are protected.</p><p><italic>If antibodies are the main effectors against influenza and CD8 T cells are not sufficient or required for the protection against influenza challenge, this should be clarified by the authors</italic>.</p><p>CD8 T cells are indeed required for the protection against the heterotypic influenza challenge as presented here in <xref ref-type="fig" rid="fig5">Figure 5</xref>. However, antibodies are essential for the protection against influenza infection such as <xref ref-type="fig" rid="fig2 fig3 fig4">Figure 2 to 4</xref>. In fact a recent paper in Nature Medicine (ref 6) shows that mice with B cell–specific deletion of Atg7 (B/<italic>Atg7</italic><sup>-/-</sup> mice) showed normal primary antibody responses after immunization against influenza but failed to generate protective secondary antibody responses when challenged with influenza virus, resulting in high viral loads, widespread lung destruction and increased fatality. We have now made the requirement of CD8+ T cells clear in the Discussion.</p><p><italic>In addition, the authors should more fully discuss the data on the role of human CD8 T cell responses in protection against influenza challenge after vaccines. The argument that the findings in mice may have relevance to diminished vaccine responses to flu vaccines in elderly relies on the relevance of CD8 T cell memory responses to vaccine efficacy. The final paragraph mentions briefly that CD8 T cell responses are an important correlate of vaccine protection; however, this should be discussed more extensively in light of the lack of in-depth experimental evidence in this study in the mouse for a role of memory CD8 T cells in protection against disease caused by viral challenge.</italic></p><p>We fully agree with the reviewers and both these points (role of CD8 T cells in humans and the lack of depth knowledge about how mice are protected in this mouse model) have now been added to the Discussion, We fully agree with the reviewers and both these points (role of CD8 T cells in humans and the lack of depth knowledge about how mice are protected in this mouse model) have now been added to the Discussion.</p><p><italic>3) T/Atg7-/- mice show defects in T cell development and a reduction of CD4 and CD8 T cells in the spleen and lymph nodes (J. Immunol. 182:4046-4055 and</italic> <xref ref-type="fig" rid="fig2"><italic>Figure 2</italic></xref><italic>). In</italic> <xref ref-type="fig" rid="fig1"><italic>Figure 1</italic></xref><italic>, the authors should show the percentage and total numbers of antigen-specific CD8 T cells among all cells in the lung, not just among gated CD8 T cells. The authors conclude that there is “normal expansion of the antigen-specific effector CD8+ T cell compartment”. However, a general lymphopenia in CD4 and CD8 T cells in these mice should be clearly presented in</italic> <xref ref-type="fig" rid="fig1"><italic>Figure 1</italic></xref><italic>, although a comparable fold of CD8 expansion was observed</italic>.</p><p>We have now swapped <xref ref-type="fig" rid="fig1 fig2">Figure 1 and 2</xref> to emphasize the lymphopenia in naïve mice at the start of the manuscript. In <xref ref-type="fig" rid="fig1">Figure 1</xref> we clearly presented the lymphopenia both as proportions (in blood and LNs) and in absolute terms (in blood over time) for CD4+ and CD8+ T cells. We agree though that this was left out of <xref ref-type="fig" rid="fig2">Figure 2</xref> where the antigen–specific response was described. So, we have now added absolute counts for influenza-challenged mice to <xref ref-type="fig" rid="fig2s1">Figure 2–figure supplement 1a</xref>). The relevant reference from the Journal of Immunology has been added to the Results section.</p><p><italic>4) The reviewers feel that the interpretation of data in</italic> <xref ref-type="fig" rid="fig5"><italic>Figure 5</italic></xref> <italic>is complicated by lymphopenia in T/Atg7-/- mice as well as autophagy deficiency in the CD4 T cell compartment. This point needs to be addressed in the authors' response</italic>.</p><p>We agree with the reviewers that this remains a limitation of our study. Unfortunately, in the time frame, we could not generate BM chimera, vaccinate and challenge them. This point was therefore discussed as a limitation of our study at the end of the Results section of the description of <xref ref-type="fig" rid="fig5">Figure 5</xref>.</p><p><italic>5) In</italic> <xref ref-type="fig" rid="fig6"><italic>Figure 6</italic></xref><italic>, the authors should consider immunodeficiency in aged CD4 T cells (Wakikawa et al., Exp Gerontol 34, 231-242; Cheung et al., Exp Gerontol 18, 451-460). Spermidine would target all cells types in vivo, including CD4+ T helper cells. The experiments performed do not distinguish between these possibilities</italic>.</p><p>The reviewers are right in pointing out that spermidine also targets CD4+ T helper cells. In <xref ref-type="fig" rid="fig6">Figure 6g</xref> we only show the effect on antigen-specific CD8+ T cells as this is our focus in the KO model. In future experiments we plan to address if CD4 T helper cells are similarly affected by the loss of autophagy and can be modulated by spermidine treatment. We have added this to the Discussion.</p><p><italic>6) The knockout of essential genes from the autophagic cascade entails a major cellular dysfunction due to accumulating, dysfunctional mitochondria (and perhaps other organelles and protein aggregates). Hence, the phenotype of Atg7 knockout cells is not just the result of autophagy inhibition but rather that of accumulating damage. This problem should be discussed</italic>.</p><p>We added a paragraph to the Discussion that deals with the problem of accumulation of damage as we, like the reviewer, think is a major part of why autophagy is important to keep the aged organism young.</p><p><italic>It would be even better to include some of the data that are mentioned as “not shown” with regard to the phenotype of mice lacking Atg5 in T cells</italic>.</p><p>The phenotype as far as the naïve mouse is concerned has been verified using the more general vav promotor and another essential autophagy gene Atg5. As shown in the graphs below, Vav-Atg5-/- are both lymphopenic and accumulate the “virtual” memory compartment, which is identical to the phenotype observed in Vav-Atg7-/-. In fact all changes (including maturation of red blood cells, hematopoietic stem cells failure and myeloid bias, <xref ref-type="bibr" rid="bib31">Mortensen et al 2010</xref> and <xref ref-type="bibr" rid="bib55">2011</xref>) have been reproduced in this Atg5 mouse model, further confirming that changes are not Atg7 specific. We have added the most relevant data of <xref ref-type="fig" rid="fig1s2">Figure 1–figure supplements 2a–b</xref>. Furthermore T cell lymphopenia in the absence of Atg5 has previously been shown by fetal liver chimera generated by Pua at al (JExp Med 2007), cited in the original manuscript.</p><p><italic>7) The authors only use one autophagy inducer, spermidine, in their experiments. Hence, it is still possible that autophagy would be only one of the spermidine effects (knowing that spermidine that might mediate additional metabolic effects on polyamine metabolism) that accounts for the improvement of CD8 memory T cells. The paper would be more complete, if the authors used other autophagy inducers (such as resveratrol, fasting etc.). At a minimum, the current limitation should be discussed</italic>.</p><p>We tested no other autophagy inducers, although we appreciate the reviewer’s comments and the wise choice of alternative treatments. This study is merely a proof of concept that memory T cell responses can be improved by autophagy modulation. We would expect that either resveratrol and fasting would be able to achieve similar results and have added these ideas to the Discussion.</p><p><italic>8) Autophagic flux is only measured by the 'BDS feature', namely the co-localization of a lysosomal marker and LC3, measured by means of the Amnis Imagestream system. It is important to confirm these measurements (and in particular the efficiency of the ATG7 knockout) by immunoblot detection of the LC3 lipidation in the absence and presence of autophagy inhibitors</italic>.</p><p>We have now performed the LC3 blot on purified CD8+ T cells from T-Atg7-/- and wild type mice and have added this blot to <xref ref-type="fig" rid="fig1s1">Figure 1–figure supplement 1b</xref> and d. It shows reduced LC3-II to LC3-I ratio in Atg7-/- T cells both in the absence and presence of the autophagy flux inhibitor Bafilomycin. We have also added a blot showing reduced expression of Atg7 in purified Atg7-/- CD8+ T cells.</p><p><italic>9) The controls in</italic> <xref ref-type="fig" rid="fig6"><italic>Figure 6d and 6e</italic></xref> <italic>look very different with regard to the proportion of LC3-I and LC3-II. This could be due to several different factors, including differences in basal autophagy conditions (density, nutritional factors) or sample processing (i.e. freeze-thawing samples will result in degradation of LC3-I which is more labile). The discrepancy in experimental conditions that results in such a notable discrepancy in the proportion of LC3-I and LC3-II in controls of the same cell type in different figures should be resolved</italic>.</p><p>The controls in <xref ref-type="fig" rid="fig6">Figure 6d and 6e</xref> look different because they are different. All samples in <xref ref-type="fig" rid="fig6">Figure 6e</xref> but not in 6d have been treated with Bafilomycin A to see the reduction of the autophagic flux with DFMO more clearly. Below is a blot without BafA showing that basal levels are a little too low to clearly show a further reduction. The low concentration of 1µm of spd only induces further LC3 lipidation when cells are treated with DFMO. However, we feel that the original blots shown were better to bring the message across.<fig id="fig7" position="float"><label>Author response image 1.</label><graphic xlink:href="elife03706f007"/></fig></p></body></sub-article></article> |