Methods for Assessing Autophagy and Autophagic Cell Death

Tasdemir, Ezgi; Galluzzi, Lorenzo; Maiuri, M. Chiara; Criollo, Alfredo; Vitale, Ilio; Hangen, Emilie; Modjtahedi, Nazanine; Kroemer, Guido

doi:10.1007/978-1-59745-157-4_3

Methods for Assessing Autophagy and Autophagic Cell Death

Ezgi Tasdemir^3,4,5,
Lorenzo Galluzzi^3,4,5,
M. Chiara Maiuri^3,4,6,
Alfredo Criollo^3,4,5,
Ilio Vitale^3,4,5,
Emilie Hangen^3,4,5,
Nazanine Modjtahedi^3,4,5 &
…
Guido Kroemer^3,4,5

Protocol

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138 Citations

Part of the book series: Methods in Molecular Biology™ ((MIMB,volume 445))

Summary

Autophagic (or type 2) cell death is characterized by the massive accumulation of autophagic vacuoles (autophagosomes) in the cytoplasm of cells that lack signs of apoptosis (type 1 cell death). Here we detail and critically assess a series of methods to promote and inhibit autophagy via pharmacological and genetic manipulations. We also review the techniques currently available to detect autophagy, including transmission electron microscopy, half-life assessments of long-lived proteins, detection of LC3 maturation/aggregation, fluorescence microscopy, and colocalization of mitochondrion- or endoplasmic reticulum–specific markers with lysosomal proteins. Massive autophagic vacuolization may cause cellular stress and represent a frustrated attempt of adaptation. In this case, cell death occurs with (or in spite of) autophagy. When cell death occurs through autophagy, on the contrary, the inhibition of the autophagic process should prevent cellular demise. Accordingly, we describe a strategy for discriminating cell death with autophagy from cell death through autophagy.

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1 Introduction

Autophagy is an evolutionarily conserved, homeostatic process that allows for the bulk degradation of long-lived proteins and organelles (1). In eukaryotic cells, the autophagic pathway is initiated when organelles and/or portions of the cytosol destined to degradation are enclosed within double-membraned vacuoles, namely autophagosomes (also known as AV, i.e., autophagic vacuoles). To promote the degradation of their luminal content, autophagosomes fuse with lysosomes, thus forming the so-called autophagolysosomes (2). It is important to note that the mere presence of AV in cells is not a proof of elevated autophagy, because a reduced turnover of AV (e.g., due to a decreased fusion of AV with lysosomes) suffices to increase their number severalfold (3). Accordingly, ultrastructural studies have to be accompanied by evidence for increased protein and/or organelle turnover to ascertain increased autophagic activity.

The execution and regulation of the autophagic program rely on several autophagy-specific genes (atg), which are characterized by a high degree of conservation among species as distant as humans and yeast (4). Some Atg proteins are directly implicated in the formation of the autophagosome. For instance, the ubiquitination of Atg5 and Atg12 by the E1-like enzymes Atg7 and Atg10 is required to form AV (5). Microtubule-associated protein light chain 3 (LC3) is the mammalian equivalent of yeast Atg8 and exists in two forms, LC3-I and -II. LC3-I is an 18-kDa polypeptide normally found in the cytosol, whereas the product of its proteolytic maturation (LC3-II, 16 kDa) resides in the autophagosomal membranes (6). Beclin-1 is the mammalian orthologue of yeast Atg6 and localizes to the trans-Golgi network, where it participates in autophagosome formation by interacting with the class III phosphatidylinositol 3-kinase (PI 3 K) human vacuolar protein sorting factor protein 34 (hVps34) (7). Importantly, the pro-autophagic interaction between hVps34 and Beclin-1 can be inhibited by the binding of the latter to the anti-apoptotic proteins Bcl-2 or Bcl-XL (8).

To ensure the turnover of old and damaged organelles, autophagy occurs constitutively at low, basal levels. However, it is also a tightly regulated adaptive mechanism that enhances cell survival under various environmental and cellular stresses, including nutrient deprivation, oxidative stress, accumulation of misfolded proteins, bacterial and viral infection, toxic stimuli, and irradiation (9, 10). One of the most efficient triggers of autophagy is starvation. In response to culture in nutrient-free media, cells degrade nonessential components to meet the cell’s energetic demand as well as to provide metabolites for vital biosynthetic reactions. In this scenario, the suppression of autophagy by chemical inhibitors or by the downregulation of essential genes (e.g., Atg5, Atg6/Beclin-1, Atg10, or Atg12) can sensitize cells to starvation-induced death (3). Autophagy inhibition sensitizes cells also to the depletion of obligatory growth (or survival) factors, which result in decreased nutrient import through the plasma membrane. This cell death occurs without massive autophagic vacuolization and is usually accompanied by the hallmarks of apoptosis, including mitochondrial membrane permeabilization (MMP) and activation of caspases (3). In some instances, the inhibition of autophagy tends to shift the cell death subroutine to more necrotic phenotypes (11).

A very different picture emerges when the late stages of AV maturation (e.g., fusion between AV and lysosomes) are inhibited. This may be accomplished by the administration of lysosomotropic bases (hydroxychloroquine or chloroquine) (3), by inhibiting the vacuolar proton pump responsible for the luminal acidification of lysosomes (with bafilomycin A1) (3), or by the knock-down of the gene encoding lysosome-associated membrane protein 2 (LAMP-2) (12). In such conditions, nutrient-starved cells massively accumulate AV in the cytoplasm, as determined by electron microscopy or by following the redistribution of the AV marker LC3 fused with a green fluorescent protein (GFP) moiety (LC3-GFP). In spite of AV accumulation, the autophagic protein turnover is inhibited (12, 13). This example illustrates the importance of using a correct combination of methods to assess AV formation, AV turnover, and cell death.

In some cases, cells manifesting massive autophagic vacuolization can be rescued from cell death by the removal of lysosomal inhibitors and/or the readdition of nutrients, despite their sometimes disastrous morphological aspect evoking textbook images of autophagic cell death. This is especially true when the still nucleus exhibits a normal aspect and mitochondria remain energized (14). However, upon prolonged nutrient starvation and inhibition of the formation of autophagolysosomes, vacuolated cells manifest the hallmarks of apoptosis (e.g., MMP, release of toxic proteins from the mitochondrial intermembrane space, and caspase activation), indicating that the point of no return that separates cellular life and death has been trespassed (15).

Autophagy has been suggested to constitute an effector mechanism of cell death, meaning that self-eating would be the first step of self-killing. Some pharmacological studies based on the use of 3-methyladenine (3-MA, a low-affinity inhibitor of hVps34) tend to support this hypothesis. For instance, it has been proposed that autophagy is required for the death of sympathetic neurons cultured in the absence of the essential nerve cell growth factor (NGF) and in the presence of caspase inhibitors (16). Similarly, TRAIL-induced autophagy accounts for the formation of hollow acini-like structures in differentiating mammary epithelial cells (17). In this model, 3-MA can provoke luminal filling when caspase-3 is inhibited by overexpression of a Bcl-Xl transgene. However, the fact that 3-MA effectively inhibits hVps34 only at concentrations $\geq$10 mM sheds major doubts on its specificity. Reportedly, inhibition of Atg genes by small interfering RNAs (siRNAs) can prevent autophagic cell death, at least in some models. For instance, siRNA-mediated downregulation of Atg6/Beclin-1 or Atg7 has been shown to reduce type 2 cell death of human U937 cells dying in response to caspase-8 inhibition (18). siRNAs targeting Atg6/Beclin-1 and Atg5 are able to prevent the autophagic cell death of Bax$^{-/-}$Bak$^{-/-}$ mouse embryonic fibroblasts succumbing to the inhibition of topoisomerase 2 by etoposide or to inhibition of tyrosine kinases by staurosporine (19). Finally, knock-down of Atg5 (or overexpression of a dominant negative variant of Atg5) can inhibit interferon-$\gamma$ (IFN-$\gamma$)–induced autophagic vacuolization and subsequent cell death (20).

The definition of the features that may be employed to identify dead cells has been the subject of a long debate. Permeabilization of the plasma membrane, disintegration of the cell, and/or phagocytosis of the corpse have been recognized as the characters that delimit the irreversibility of the process (21, 22). Several subroutines of cell death can be distinguished according to ultrastructural criteria. The best-studied modality of cell death, apoptosis (type 1 cell death), is characterized by morphological changes that include nuclear pyknosis (chromatin condensation) and karyorhexis (nuclear fragmentation). As the process advances, cells form small round bodies surrounded by membranes that contain intact cytoplasmic organelles and/or nuclear fragments. In vivo, these “apoptotic bodies” are usually engulfed by resident phagocytic cells. As mentioned above, massive autophagic vacuolization is observed in some instances of cell death, which has been named “autophagic cell death” (type 2 cell death) (22). It is an ongoing conundrum, however, in which case “autophagic cell death” is truly mediated through autophagy (meaning that its inhibition would prevent cell death) and in which case it simply occurs together with autophagy (meaning that inhibition of autophagy would affect only the morphology of the process, but not the fate of cells) (12, 23). Necrosis (type 3 cell death) is a subroutine of cell death that does not manifest the hallmarks of apoptosis or massive autophagic vacuolization. The principal feature of necrosis is a gain in cell volume (oncosis) that finally culminates in the sudden rupture of the plasma membrane, accompanied by the unorganized dismantling of swollen organelles (11).

In spite of the large amount of published data, the methods that are currently available for the detection of autophagy are affected by numerous intrinsic pitfalls. The most reliable and conventional technique to visualize autophagic vacuolization is transmission electron microscopy. Biochemical methods and techniques designed to measure the aggregation of autophagosome markers allow for the monitoring of autophagy, yet are afflicted by relevant problems. Here, we will provide an overview of routine methods for the assessment of autophagy and autophagic cell death.

2 Materials

2.1 Common Materials

2.1.1 Disposables

1.
1.5-mL microcentrifuge tubes (Eppendorf, Hamburg, Germany).100 $\times$ 20 culture dishes (Corning Inc. Life Sciences, Acton, MA).
2.
12-mm $\O$ cover slips (Menzer-Gläser GmbH, Braunschweig, Germany)—sterilized by incubation for 15–30 min in 100% ethanol (Carlo Erba Reagents, Milan, Italy).
3.
15- and 50-mL conical centrifuge tubes (BD Falcon, San Jose, CA). 175-cm$^{2}$ flasks for cell culture (BD Falcon).
4.
5-mL, 12 $\times$ 75 mm FACS tubes (BD Falcon).
5.
6-, 12-, 24-well plates for cell culture (Corning Inc. Life Sciences).
6.
76 $\times$ 26 mm slides for fluorescence microscopy (Carl Roth GmbH, Karlsruhe, Germany).

2.1.2 Solutions

1.
Growth medium for HeLa cells: Dulbecco’s modified Eagle’s medium (DMEM) containing 4.5 g/L glucose, 4 mM L-glutamine and 110 mg/L sodium pyruvate (Gibco-Invitrogen, Carlsbad, CA) supplemented with 100 mM HEPES buffer (Gibco-Invitrogen) and 10% fetal bovine serum (FBS, from PAA Laboratories GmbH, Pasching, Austria).
2.
PBS (1X): 137 mM NaCl, 2.7 mM KCl, 4.3 mM Na$_2$HPO$_4$, 1.4 mM KH$_2$PO$_4$ in deionized water (dH$_2$O), adjust pH to 7.4 with 2 N NaOH.
3.
Trypsin/ethylenediaminetetraacetic acid (EDTA): 0.25% trypsin, 0.38 g/L (1 mM) EDTA${\bullet}$4Na in Hank’s balanced salt solution (HBSS) (Gibco-Invitrogen).

2.2 Experimental Modulation of Autophagy

1.
Autophagy inducers are listed in Table 1.
Table 1 Autophagy Inducers
Full size table
2.
Autophagy inhibitors are listed in Table 2.
Table 2 Autophagy inhibitors
Full size table
3.
siRNAs used for the inhibition of the autophagic pathway in human cell lines are reported in Table 3.
Table 3 siRNAs Used to Knock Down Genes Involved in utophagy in Human Cell Lines
Full size table
4.
Oligofectamine™ transfection reagent (Oligofectamine™ from Gibco-Invitrogen).
5.
Opti-MEM^® reduced serum medium (Opti-MEM^®), with Glutamax™ and phenol red (Gibco-Invitrogen).

2.3 Measuring Autophagy

1.
3 MM^® Whatmann filter paper.
2.
40% acrylamide:N-N’-methylenediacrylamide (29/1) solution (Bio-Rad, Hercules, CA).
3.
6-mm-thick, 10.5 $\times$ 11 cm foam sponges (GE Healthcare Life Sciences Sciences, Uppsala, Sweden).
4.
Ammonium persulfate (APS), stock solution at 10% (w/v) in dH$_2$O (stored at 4$^\circ$C) (see also Note 37).
5.
Antibodies employed in immunoblotting techniques are listed in Table 4.
Table 4 Antibodies for Immunoblotting
Full size table
6.
Blocking buffer: 0.1% Tween-20 (Sigma-Aldrich) and 5% (w/v) nonfat powdered milk (commonly found in food stores) in PBS. Storage at 4$^\circ$C should not exceed one week, unless 0.02% sodium azide is added as preservative.
7.
Bovine serum albumin (BSA, from Sigma-Aldrich), stock solution 10 mg/mL in dH$_2$O, stored at 4$^\circ$C.
8.
Chemiluminescent substrate: SuperSignal West Femto Maximum Sensitivity Substrate (Pierce Biotechnology, Rockford, IL).
9.
Chemiluminescent substrate: SuperSignal West Pico Chemiluminescent Substrate (Pierce Biotechnology).
10.
DC protein assay (Bio-Rad).
11.
Dehybridizing buffer: 570 $\mu$L of 1 N acetic acid solution (Sigma-Aldrich) + 100 mL dH$_2$O.
12.
Electrophoresis apparatus: Mini-PROTEAN 3 electrophoresis cell system (Bio-Rad), powered by a PowerPac HC power supply (Bio-Rad).
13.
Epon^® epoxy resin (Miller-Stephenson, Danbury, CT).
14.
Film processor: Curix 60 table-top film processor (Agfa, Mortsel, Belgium).
15.
Fixative solution: 4% paraformaldehyde (PFA, from Sigma-Aldrich) + 0.19% picric acid (Sigma-Aldrich) in PBS. For 10 mL: dissolve 0.4 g of PFA in 500 $\mu$L of dH$_2$O, add one drop of 2 N NaOH, heat to 65–70$^\circ$C until solution clears, add 9.5 mL of PBS, let cool to room temperature (RT), add 38 $\mu$L of saturated picric acid (Sigma-Aldrich), and move the fixative solution to ice bath (see also Notes 13–15).
16.
Fluorescence microscope: IRE2 microscope equipped with a DC300F camera (Leica Microsystems GmbH, Wetzlar, Germany).
17.
Fluorescent dyes used for the detection of autophagy are listed in Table 5.
Table 5 Fluorescent Probes, Labels, and Proteins for the Detection of Autophagy and Autophagic Cell Death
Full size table
18.
Fluoromount-G™ mounting medium (Southern Biotech, Birmingham, AL).
19.
Grade I glutaraldehyde solution, 25% in dH$_2$O, specially purified for use as an electron microscopy fixative (Sigma-Aldrich).
20.
Isopropyl alcohol (isopropanol, CAS No. 67-63-0, from Carlo Erba Reagents).
21.
l-[U-$^{14}$C]-Valine, aqueous solution containing 2% ethanol, sterilized (GE Healthcare Life).
22.
Lead citrate: dissolve 0.1 g of lead(II) citrate tribasic trihydrate (Sigma-Aldrich) in 100 mL dH$_2$O; add dropwise 10 N NaOH (Sigma-Aldrich) until solution becomes clear; store at RT.
23.
Lipofectamine™ 2000 transfection reagent (Lipofectamine™, from Gibco-Invitrogen).
24.
Loading buffer (5X): 313 mM Tris-HCl (pH 6.8), 10 mM EDTA, 500 mM dl-dithiothreitol (DTT), 10% (w/v) sodium dodecylsulfate (SDS), 50% (v/v) glycerol, 0.05% (w/v) bromophenol blue in dH$_2$O (or commercially available from Fermentas, Ontario, Canada).
25.
L-Valine (Sigma-Aldrich), stock solution in dH$_2$O, at the concentration of 100 mM (stored at 4$^\circ$C).
26.
Lysis buffer: CelLytic™ M (Sigma-Aldrich).
27.
Migration buffer (1X): 100 mL 10X Tris/Glycine/SDS buffer (Bio-Rad) + 900 mL dH$_2$O.
28.
Mini-PROTEAN 3 combs (5-, 9- 10- or 15- wells, from Bio-Rad) (see also Note 46).
29.
N,N,N’,N’-Tetra-methyl-ethylenediamine (TEMED, from Bio-Rad).
30.
Nitrocellulose membrane roll (Bio-Rad).
31.
Osmium tetroxide solution for electron microscopy, 2% in dH$_2$O.
32.
Photographic films: 18 $\times$ 24 Amersham Hyperfilm™ ECL™ (GE Healthcare Life Sciences).
33.
Ponceau S staining solution: 0.1% (w/v) Ponceau S and 5.0% (w/v) acetic acid (Sigma-Aldrich) in dH$_2$O (or commercially available from Sigma-Aldrich).
34.
Protein molecular weight (MW) markers: Precision Plus Protein™ Standard, Dual Color (Bio-Rad).
35.
Rinsing buffer: 0.1% Tween-20 in PBS. Storage at RT.
36.
Running gel buffer: 1.5 M Tris buffer (pH 8.8, from Bio-Rad).
37.
Sörensen phosphate buffer (1X, 0.1 M, ph 7.3): 23 mL of 0.2 M NaH$_2$PO$_4$ + 77 mL of 0.2 M Na$_2$HPO$_4$ + 100 mL dH$_2$O.
38.
Stacking gel buffer: 0.5 M Tris buffer (pH 6.8).
39.
Transfer buffer (1X): 100 mL 10X Tris/glycine buffer (Bio-Rad) + 200 mL ethanol + 900 mL dH$_2$O.
40.
Transfer cassette: Mini Trans-Blot Cell (Bio-Rad).
41.
Transmission electron microscope: Tecnai G2 Spirit (FEI, Eindhoven, The Netherlands).
42.
Transparent plastic film (Saran, as used for food preservation).
43.
Trichloroacetic acid (TCA), stock solution 10% (w/v) in dH$_2$O (stored at RT).
44.
Uranyl acetate: dissolve 10 g of uranyl acetate dihydrate in 100 mL of methanol (Carlo Erba Reagents); shake or vortex until complete dissolution; store at $-$20$^\circ$C.

2.4 Measurement of the Colocalization of Cytoplasmic Organelles with Autophagic Vacuoles

1.
0.1% (w/v) SDS in PBS.
2.
10% FBS (PAA Laboratories GmbH) in PBS.
3.
Antibodies employed for immunofluorescence microscopy assessments are listed in Table 6.
Table 6 Antibodies for Immunofluorescence Microscopy
Full size table
4.
BSA buffer: 3 mg/mL BSA (Sigma-Aldrich) in PBS.
5.
Confocal microscope: Zeiss LSM 510 (Carl Zeiss AG, Oberkochen, Germany).
6.
Fluorochromes used to measure the colocalization of cytoplasmic organelles with autophagic vacuoles are listed in Table 5.

2.5 Measuring Cell Death–Related Parameters

1.
Annexin V-FITC kit for the detection of phosphatidylserine externalization (Miltenyi Biotec, Bergisch Gladbach, Germany).
2.
Carbonyl cyanide m-chlorophenylhydrazone (CCCP, from Sigma-Aldrich), stock solution in ethanol, 10 mM, stored at $-$20$^\circ$C.
3.
Cytofluorometer: FACScan (Becton Dickinson, San Jose, CA) equipped with an argon ion laser emitting at 488 nm.
4.
Fluorescent probes used to measure cell death–related parameters are listed in Table 5.

3 Methods

Cells that activate the autophagic program, manifest in the accumulation of double-membraned autophagic vesicles in the cytoplasm. These vacuoles can be classified according to their electron density, as assessed by transmission electron microscopy, into early and late AV. Early AV (AV1) contain cytoplasmic portions and/or organelles and exhibit an electron density equivalent to the cytoplasm (AV1). Late or degradative AV (AV2) show evidence of partially digested luminal content and are characterized by increased electron density.

The mere presence of AV in the cytoplasm does not necessarily indicate an increased level of autophagy, since a reduction of the fusion between autophagosomes and lysosomes suffices to promote AV accumulation. Thus, quantitative methods for the detection of cytoplasmic protein turnover should be employed in addition to AV monitoring, in order to verify augmented levels of autophagy.

3.1 Experimental Modulation of Autophagy

Given its importance in cellular homeostasis and response to stress, it is not surprising that autophagy is a precisely controlled process that is regulated by several (cross-talking) signaling pathways (24, 25). In this context, a major role is played by the mammalian target of rapamycin (mTOR) protein, a serine/threonine kinase implicated in the control of several cellular functions (26, 27). mTOR integrates numerous inputs (including signals from growth factors, insulin, cellular stressors as well as those reporting the availability of nutrients) to stimulate protein synthesis via the phosphorylation of key translation regulators like ribosomal protein S6 kinase (p70SK6) and eukaryotic initiation factor 4E binding protein 1 (eIF4EBP1) (28). Accordingly, mTOR is a major gatekeeper and inhibitor of autophagy, and may exert its anti-autophagic effects either by promoting protein synthesis or by a direct inhibition of crucial atg proteins. Other important regulators of autophagy include class I and class III PI 3 Ks. The increase in class I PI 3 K products (like phosphatidylinositol 3,4-bisphospate, i.e., PIP2, and phosphatidylinositol 3,4,5-trisphospate, i.e., PIP3), caused by feeding the cells with synthetic lipids or via by stimulating the interleukin-13 receptor, has been reported to inhibit macroautophagy (29). Conversely, class III PI 3 K (hVps34) activates autophagy by interacting with Beclin-1 and plays a crucial role at early stages of autophagosome formation (30).

3.1.1 Induction of Autophagy

In vitro, several strategies may be employed to induce autophagy (Fig. 1). These include: the inhibition of mTOR (e.g., by starvation, by inhibition of the upstream class I PI 3 K-mediated signaling, or by administration of the mTOR specific inhibitor rapamycin) (27, 31); the inhibition of inositol monophosphatase (IMPase), resulting in a reduction of free inositol and inositol-1,4,5-triphosphate (IP3) levels (e.g. with lithium, L-690,330 or carbamazepine) (32); the blockade of the IP3 receptor (IP3R) of the endoplasmic reticulum (ER) (e.g., with xestospongin B and C) (33); the induction of ER stress via the activation of the unfolded protein response (e.g., with tunicamycin, thapsigargin or brefeldin A) (34); or the induction of oxidative stress (e.g. with H$_2$O$_2$) (35).

figure 3_1_978-1-59745-157-4 — **Fig. 1**

1.
Wild-type HeLa cells are cultured in appropriate growth medium and passaged when approaching to confluence with trypsin/EDTA to provide new maintenance cultures in 175-cm$^{2 }$flasks and experimental cultures. The latter are performed either in 12-well plates (100 $\times$ 10$^{3 }$cells in 1 mL of growth medium) or in 24-well plates in which sterile cover slides have been previously deposited (25 $\times$ 10$^{3}$ cells in 0.5 mL of growth medium).
2.
Twelve to 24 h after plating, induction of autophagy is performed. To this aim, growth medium is removed by aspiration and cells are washed twice with 1 or 2 mL (in 24- or 12-well plates, respectively) sterile PBS. Then, autophagy inducers (see Table 1) are administered in growth medium at the appropriate concentration (see Note 1). Alternatively, growth medium is substituted with 0.5–1 mL of Earle’s balanced salt solution (EBSS, nutrient-free [NF] medium). Finally, cells are incubated at 37$^\circ$C in 5% CO$_2$ atmosphere for a variable time frame, depending on the experimental setting (see Note 2).
3.
Finally, autophagic activity and cell death can be monitored as described below (see Subheading 3.2.).

3.1.2 Inhibition of Autophagy

The cascade of events leading to autophagy can be suppressed at various steps (36) (Fig. 1), which include: (1) the formation of autophagosomes; (2) the fusion between autophagosomes and lysosomes (3, 13); and (3) the final, lysosomal degradation phase (15, 37). The inhibition of the autophagic process can be achieved by different means, notably (1) chemical inhibitors (see Table 2) (3, 13, 15); (2) transfection-enforced overexpression of inhibitory proteins (8); and (3) siRNAs targeting essential atg gene products (see Table 3) (3, 12, 30).

3-Methyladenine (3-MA) prevents autophagy at the sequestration step by inhibiting the class III PI 3 K hVps34 (29). However, the results of experiments in which 3-MA is employed as an autophagy inhibitor should be evaluated cautiously due to the fact that this compound affects many other signaling pathways (by interacting with kinases such as class I PI 3 K; c-Jun N-terminal kinase, i.e., JNK; and mitogen-activated protein kinases, i.e., MAPK) (29, 38). Moreover, at the concentration of 10 mM (representing the minimal concentration required to block autophagy), 3-MA also inhibits MMP and other catabolic pathways (39).

Hydroxychloroquine is a lysosomotropic amine, which is clinically used for the treatment of malaria, rheumatoid arthritis, and systemic lupus erythematosus. As do other immunosuppressant drugs, hydroxychloroquine exhibits cytotoxic properties. This agent induces the alkalinization of the lysosomal lumen, thereby inhibiting the fusion between lysosomes with autophagosomes and interfering with the action of lysosomal hydrolases, which function at an acidic pH. At higher concentrations, hydroxychloroquine induces lysosomal membrane permeabilization (LMP), resulting in the potentially lethal leakage of catabolic hydrolases into the cytosol (15).

Bafilomycin A1 is a specific inhibitor of the vacuolar H$^{+}$-ATPase, the proton pump that acidifies the lysosomal lumen (37, 40). Similarly to hydroxychloroquine, bafilomycin A1 prevents the activation of lysosomal hydrolases and interferes with the fusion between autophagosomes and lysosomes (13), presumably because lysosomal acidification is required for this step.

The intrinsic disadvantage of chemical inhibitors is that they can exert several unwanted side effects due to the fact that none of the currently available molecules is truly specific for autophagy. The siRNA-mediated depletion of proteins essential for the autophagic pathway is characterized by a high degree of specificity. siRNAs targeting hVps34, Beclin-1, Atg5, Atg10, Atg12, and LAMP-2 are now widely employed to suppress autophagy (see Table 3) (3, 12, 30).

3.1.2.1 Chemical Inhibition of Autophagy

1.
HeLa cells are maintained in culture and plated as described above (see Subheading 3.1.1.).
2.
Chemical inhibition of autophagy is performed 12–24 h after plating. As for the induction of autophagy, growth medium is removed, cells are washed twice with sterile PBS, then autophagy inhibitors (see Table 2) are administered at the appropriate concentration (see Note 1), either in growth medium (negative control condition) or in NF medium (providing a positive control for autophagy induction). Thereafter, cells are cultured at 37$^\circ$C in a 5% CO$_2$ incubator.
3.
After a time frame varying according to the specific experimental aim (see Note 2), autophagic activity can be measured as described below (see Subheading 3.2.).

3.1.2.2 siRNA-Mediated Inhibition of Autophagy

1.
HeLa cells are seeded in 6-well plates (approximately, 200 10$^{3}$ cells in 3 mL of growth medium), in order to obtain confluence levels around 60–70% on the next day.
2.
If cells have reached appropriate confluence, transfection of siRNA is performed 12–24 h after plating (see Note 3). For the delivery of siRNAs to adherent cells, liposome-based transfection is employed. To this aim, 200 nmol of siRNA (final concentration in wells = 100 nM) are diluted in 180 $\mu$L of Opti-MEM^® and 4 $\mu$L of Oligofectamine™ transfection reagent (Oligofectamine™) are gently mixed with 16 $\mu$L of Opti-MEM^®. After a first incubation of 5–10 min, the diluted siRNAs and the diluted Oligofectamine™ solution are combined, gently mixed, and incubated for additional 20 min to allow for the formation of Oligofectamine™:siRNA transfection complexes (see Notes 4 and 5).
3.
Before the addition of the complexes to the cells, growth medium is replaced by 1.8 mL of fresh medium without serum (see Notes 6 and 7). Then, 200 $\mu$L of the solution containing Oligofectamine™:siRNA complexes are added to each well, and plates are incubated at 37$^\circ$C in 5% CO$_2$ atmosphere for 4 h.
4.
Four hours after transfection, 220 $\mu$L of FBS should be added to each well to restore the final concentration of 10% (as in complete growth medium).
5.
Twelve to 24 h after transfection, cells can be trypsinized (500 $\mu$L of Trypsin/EDTA per well) and seeded in 12- or 24-well plates, according to the specific experimental settings (see Notes 8–10).
6.
After the time required for the siRNA-mediated downregulation of the target protein (see Note 8), the desired stimuli can be administered to the cells in which the autophagic pathway has been suppressed by genetic techniques.
7.
Finally, following another incubation at 37$^\circ$C in 5% CO$_2$ atmosphere (the duration depends again on the specific experimental conditions), autophagy can be assessed as described below (see Subheading 3.2.).

3.2 Measuring Autophagy

At present, multiple techniques are available for the quantification of autophagy and the identification of AV. Table 7 summarizes these methods, by providing also the main advantages and drawbacks of each.

Table 7 Methods for the Detection of Autophagy 7

Full size table

3.2.1 Electron Microscopy

1.
2 $\times$ 10$^{6 }$cells are cultured in 100 $\times$ 20 mm culture dishes.
2.
After the treatment with the desired stimuli, cells are fixed for 1 h at 4$^\circ$C in 1.6% glutaraldehyde in 0.1 M Sörensen phosphate buffer (pH 7.3) and washed once with PBS.
3.
Cells are then re-fixed in aqueous 2% osmium tetroxide and finally embedded in Epon^® epoxy resin, until imaging.
4.
Examination is performed at 80 kV under a transmission electron microscope, on ultrathin sections (80 nm) stained with 0.1% lead citrate and 10% uranyl acetate.

Once the images are obtained, the number of type I and type II AV can be quantified on a per-cell basis (see Fig. 2). In addition, AV volume can be evaluated by morphometric methods.

figure 3_2_978-1-59745-157-4 — **Fig. 2**

3.2.2 LC3-GFP Aggregation

1.
HeLa cells (25 $\times$ 10$^{3}$ in 0.5 mL growth medium) are seeded in 24-well plates in which sterile cover slips have been previously deposited.
2.
Twelve to 24 h later, cells are transfected with a plasmid coding for the autophagosome marker LC3 fused with green fluorescence protein (GFP) (6), according to the following protocol. 4 $\mu$g of plasmid are diluted in 200 $\mu$L of Opti-MEM^®, at the same time as 5 $\mu$L of Lipofectamine™ are gently mixed with 200 $\mu$L Opti-MEM^®. After a first incubation of 5–10 min, the diluted plasmid solution and diluted Lipofectamine™ solution are gently mixed and incubated for another 20 min to promote the formation of Lipofectamine™:plasmid complexes (see Notes 4 and 5).
3.
Thereafter, 30 $\mu$L of solution containing the Lipofectamine™:plasmid complexes are added to each well, in which medium had been previously replaced with 500 $\mu$L of serum-free growth medium. Plates are then incubated at 37$^\circ$C in 5% CO$_2$ atmosphere for 4 h before 60 $\mu$L of FBS are added to restore the final FBS concentration of 10% (as in complete growth medium) (see Note 6).
4.
Cells are cultured for 24 h, or until they start to express the LC3-GFP fusion protein, prior to treatment with the desired stimuli (see Notes 11 and 12).
5.
At the end of stimulation, growth medium is removed, cells are washed twice with PBS and fixed in 400 $\mu$L of fixative solution for 30 min at RT (see Notes 13–16).
6.
Fixative solution is removed, cells are washed three times with PBS, and nuclear counterstaining is performed by the addition of 2 $\mu$ M Hoechst 33342 in PBS (200 $\mu$L per well) for 10 min (see Note 17).
7.
The cover slips are mounted onto slides using Fluoromount-G™ mounting medium prior to examination in a fluorescence microscope. For the visualization of LC3-GFP, cells should be examined using an oil immersion objective (50–65$\times$ magnification). Suitable excitation and emission filters should be selected according to the following absorption and emission peaks: 488/507 nm for LC3-GFP and 352/461 nm for Hoechst 33342 (see Note 18). The LC3-GFP fusion protein redistributes from a diffuse (cytoplasmic and nuclear) to a vacuolar, punctuate (exclusively cytoplasmic) pattern when AV are formed (see Fig. 3 and Note 19)
Fig. 3
Detection of autophagy by the LC3-GFP aggregation technique. HeLa cells transfected with a plasmid encoding the LC3-GFP fusion protein (as described in Subheading 3.2.2.) were left untreated (Control) or treated with 2.5 $\mu$ M tunicamycin for 6 h (Autophagy). Thereafter, cells were processed for fluorescence microscopy examination, as detailed in Subheading 3.2.2. In untreated cells LC3-GFP exhibits a diffuse cytoplasmic signal. When autophagy is induced, LC3-GFP chimeric proteins aggregate in autophagic vacuoles, leading to a punctuate cytoplasmic staining. As confirmed by Hoechst 33342 counterstaining, LC3-GFP is not found in the nucleus.
Full size image

3.2.3 Bulk Degradation of Long-Lived Proteins

1.
HeLa cells (400 $\times$ 10$^{3}$ in 2.5 mL growth medium) are plated in 6-well plates.
2.
Forty-eight hours later, growth medium is replaced by 0.2 $\mu$Ci/mL L-[U-$^{14}$C]valine (see Note 20) in complete medium. Plates are then incubated at 37$^\circ$C in 5% CO$_2$ atmosphere for 24 h.
3.
At the end of the radiolabeling period, cells are washed three times with PBS (to remove unincorporated radioisotopes), and incubated with 10 mM unlabeled L-valine in complete growth medium (2–2.5 mL) for 1 h (prechase period) (see Note 21).
4.
After the prechase step, the culture medium is again replaced by fresh medium containing 10 mM unlabeled valine in the presence or absence of an autophagy inhibitor (e.g., 3-MA) (see Note 22). Cells are then incubated with the desired stimuli for 4–12 h (chase period).
5.
After the chase step, supernatant is collected and proteins from the medium and from cells are precipitated separately with 10% (w/v) TCA (overnight, 4$^\circ$C).
6.
Thereafter, TCA-precipitated fractions are centrifuged (600g, 20 min, RT) and redissolved in 1 mL of 0.2 N NaOH. Finally, radioactivity is determined by liquid scintillation counting.
7.
The rate of degradation of long-lived proteins can be calculated by determining the ratio of TCA-precipitated radioactivity recovered from the medium to the sum of TCA-precipitated radioactivity from the medium and cells:
$$\% {\rm Proteolysis (h^{-1}) = R_m / (R_c + R_m)}$$

where R$_{\rm m}$ is TCA-precipitated radioactivity from medium and R$_{\rm c}$ is TCA-precipitated radioactivity from cells.

3.2.4 Monodansylcadaverine (MDC) Staining of Autophagic Vacuoles

1.
HeLa cells (25 $\times$ 10$^{3}$ in 0.5 mL of growth medium) are seeded in 24-well plates where sterile cover slips have been previously dropped.
2.
Following the desired stimuli for the induction or inhibition of autophagy, AV are labeled with 50 $\mu$ M MDC in growth medium (0.5 mL) for 30 min at 37$^\circ$C.
3.
Then, cells are washed three times with PBS and fixed in 400 $\mu$L of 4% PFA (see Notes 13 and 14) for 30 min at RT. After fixation, PFA is removed and cells are washed once with PBS. Finally, cover slips are mounted as described above (see Subheading 3.2.2., step 7).
4.
As for LC3-GFP staining, the assessment of AV is performed by fluorescence microscopy, at magnifications of 50–65$\times$. Usually, MDC is observed with a 335(380)/525 nm (excitation/emission) filter set (see Note 23). The percentage of cells with punctuate structures should be assessed among a sample of statistical relevance (see Notes 19 and 24).

3.2.5 CellTracker™ Green (CMFDA) Staining

1.
25 $\times$ 10$^{3}$ HeLa cells are cultured in 24-well plates with sterile coverslips (0.5 mL growth medium).
2.
After the stimulation or inhibition of autophagy, cells are stained with 1 $\mu$ M CMFDA at 37$^\circ$C in a 5% CO$_2$ incubator for 30 min.
3.
Then, cells can be fixed and counterstained with Hoechst 33342 as described above (see Subheading 3.2.2., steps 5–7, and Notes 13–17).
4.
CMFDA staining is routinely examined in fluorescence microscopy with a 488/525 (excitation/emission) filter set (see Note 25).
5.
With this technique, AV vacuoles appear as “holes” within a diffuse green cytoplasmic fluorescence. The percentage of cells bearing at least one discernable cytoplasmic vacuole should be determined (see Note 19).

3.2.6 LC3-I–to–LC3-II Conversion by Immunoblotting

3.2.6.1 Preparation of Samples

1.
HeLa cells (5 $\times$ 10$^{5 }$in 3 mL of growth medium) are cultured in 6-well plates.
2.
Following the desired treatments to induce or suppress autophagy, cells are collected by trypsinization, washed once in cold (4$^\circ$C) PBS and pelleted (300–500g, 5 min, 4$^\circ$C) (see Note 26).
3.
Supernatant is discarded, and pellets are lysed in 30–50 $\mu$L of lysis buffer (see Notes 27 and 28).
4.
The protein concentration of lysates is determined by means of the Bio-Rad DC Protein Assay, following the manufacturer’s instructions (see Note 29).
5.
Then, 4–6 $\mu$L of 5X loading buffer are added to 20–80 $\mu$g of proteins from each lysate, and the final volume is adjusted to 20–30 $\mu$L with dH$_2$O (see Notes 30 and 31).
6.
Finally, samples are incubated for 5 min at 100$^\circ$C. After cooling to RT, samples are ready for separation by sodium dodecylsulfate–polyacrylamide gel electrophoresis (SDS-PAGE) (see Note 32).

3.2.6.2 SDS-PAGE

1.
These instructions assume the use of a BioRad mini-PROTEAN 3 system for gel electrophoresis. The protocol is easily adaptable to other formats.
2.
A 1.0-mm-thick, 15% running gel (see Note 33) is prepared by mixing 1.8 mL of 40% acrylamide:N-N’-methylenediacrylamide (29/1) solution (see Note 34), 1.25 mL of running gel buffer (pH 8.8), 1.8 mL of dH$_2$O, 50 $\mu$L of 10% SDS, 10 $\mu$L of TEMED (see Notes 35 and 36), and 26 $\mu$L of 10% APS (see Notes 37 and 38). Immediately after the addition of APS (to avoid polymerization in tube; see Notes 39 and 40), gel should be poured into preassembled glasses up to approximately two thirds of their height (see Notes 41 and 42).
3.
Quickly following, overlay gel with isopropanol (see Note 43) to ensure a flat surface and exclude air.
4.
After the polymerization (see Note 44), isopropanol is poured off and the top of the gel is washed with dH$_2$O.
5.
The stacking gel (see Note 45) is prepared by mixing 250 $\mu$L of 40% acrylamide/bisacrylamide (29/1) solution (see Note 34), 625 $\mu$L of stacking gel buffer (pH 6.8), 1.6 mL of dH$_2$O, 50 $\mu$L of 10% SDS, 9 $\mu$L of TEMED (see Notes 35 and 36), and 15 $\mu$L of APS (see Notes 37 and 38). Rapidly following the addition of APS, the stacking gel is poured on top of the polymerized running gel and the comb for the creation of wells is inserted (see Notes 38 and 46).
6.
Once the stacking gel is polymerized, the comb is gently removed and the gel unit is moved to the cell for separation. Upper and lower chambers of the gel unit are filled with migration buffer (1X) (see Note 47), which is used also to gently wash sample wells.
7.
At this stage, each well is loaded with a sample prepared as described above (see Subheading 3.2.6.1.). One well is reserved for protein MW markers (see Note 48).
8.
Finally, the gel unit is connected to power supply and the proteins are separated in constant-field mode (see Note 49). The run should be arrested when MW markers are separated throughout entire height of the gel (see Note 50).

3.2.6.3 Immunoblotting (Western Blotting)

1.
Two sponges and four sheets of filter paper of approximately the same size as the running gel are incubated in transfer buffer (1X) for 1–2 min at RT.
2.
One nitrocellulose membrane of approximately the same size of the running gel is activated by incubation in dH$_2$O for 5–10 min at RT, then equilibrated in transfer buffer.
3.
The gel unit is disconnected from the power supply and disassembled. The stacking gel is discarded.
4.
The transfer “sandwich” is prepared by overlaying the following components (see Note 51): one sponge, two sheets of Whatmann paper, the nitrocellulose membrane, the running gel, two sheets of Whatmann paper, one sponge.
5.
The “sandwich” is placed in a transfer cassette (see Note 52), which is subsequently inserted into the transfer apparatus. Special attention should be paid in the orientation of the cassette to ensure that the nitrocellulose membrane is placed between the gel and the anode (see Note 53).
6.
To control temperature, an ice block is placed next to the transfer cassette (see Note 54). The cell is then completely filled with transfer buffer (1X). A magnetic stir-bar is placed in the cell to help in maintaining a homogeneous solution and temperature.
7.
The power supply is activated, and the transfer is accomplished at the constant voltage of 90 V for 2 h (see Notes 55 and 56).
8.
Once the transfer is complete, the cassette is disassembled. Sponges and sheets are removed (see Note 57). The colored MW markers should be visible on the nitrocellulose membrane.
9.
The bound proteins can be visualized by a rapid incubation with Ponceau S staining solution (see Note 58). Prior to the blocking, Ponceau S should be removed by rinsing the membrane with PBS or dH$_2$O.
10.
To block unspecific binding sites, the nitrocellulose membrane is incubated on a rocking platform in 10 mL of blocking buffer (45–40 min, RT).
11.
Blocking buffer is discarded, membrane is quickly rinsed in rinsing buffer and a 1/200 dilution of rabbit anti-LC3 polyclonal antibody (in 5–10 mL of blocking buffer) is added. Incubation with the primary antibody can be performed either at RT for 2 h or at 4$^\circ$C overnight, in both cases on the rocking platform.
12.
Once the primary antibody is removed, the membrane is washed three times with rinsing buffer (5 min, RT).
13.
Upon washing, membrane is incubated for 1 h at RT (on the rocking platform) with a freshly prepared 1/5000 dilution in blocking buffer of goat anti-rabbit horseradish peroxidase (HRP)–labeled secondary antibody.
14.
Secondary antibody solution is removed and the membrane is again washed three times (5 min, RT) with rinsing buffer.
15.
The membrane is overlaid with 2 mL of chemiluminescent substrate for 1 min (see Note 59). Then, chemiluminescent substrate is removed and the membrane is moved to an x-ray film cassette.
16.
In the dark room, a photographic film (see Note 60) is exposed to the membrane for a suitable time, typically 3 or 4 min (see Notes 61 and 62).
17.
Finally, exposed films are developed in a common table-top film processor.
18.
To ensure equal loading of the samples onto the gel (thus comparability of the bands), membrane can be dehybridized from bound antibodies by incubation in dehybridizing buffer for 15 min at RT, followed by one wash in rinsing buffer and by repetition of the steps 10–17 with the following antibodies:
- Primary antibody: mouse anti-glyceraldehyde-3-phosphate dehydrogenase (GAPDH) monoclonal antibody (see Note 63), 1/10000 in 10 mL of blocking buffer, 1 h, RT.
- Secondary antibody: goat anti-mouse HRP-labeled secondary antibody, 1/5000 in 10 mL of blocking buffer, 1 h, RT.

LC3-II is the posttranslationally processed form of LC3, which localizes to autophagosomes and autophagolysosomes during autophagy(6). Under nonautophagic conditions, the AV marker LC3 appears as a single band of approximately 18 kDa, corresponding to LC3-I, the uncleaved isoform. Following the induction of autophagy, LC3-II (the cleaved isoform of LC3) can be visualized as an additional band with a slightly higher electrophoretic mobility than LC3-I (approximately 16 kDa).

3.3 Methods to Measure the Colocalization of Cytoplasmic Organelles with Autophagic Vacuoles

3.3.1 Lysosomes

Lysosomes are acidic vesicles that participate in the degradation of extracellular macromolecules that are taken up via endocytosis, micropinocytosis, or phagocytosis. Lysosome are also implicated in the catabolism of cytoplasmic components through autophagy(36). The most abundant proteins found in lysosomal membranes are the lysosome-associated membrane proteins 1 and 2 (LAMP-1 and -2), which together constitute almost 50% of all lysosomal membrane proteins. To study the fusion between AV and lysosomes, colocalization experiments can be performed according to different staining techniques.

3.3.1.1 LAMP-2 Immunofluorescence Staining

1.
HeLa cells are cultured in 24-well plates, transfected with the LC3-GFP plasmid and treated as described above (see Subheading 3.2.2., steps 1–4, and Notes 4–6, 11, and 12).
2.
After the desired stimulation, growth medium is discarded, cells are washed twice with PBS, and incubated with fixative solution for 30 min at RT. Thereafter, cells are rinsed an additional three times with PBS (see Notes 13–16).
3.
Incubation for 10 min (RT) in 0.1% SDS is used to permeabilize cells. Then, cells are rinsed three times with PBS and nonspecific binding sites are blocked by incubating the samples for 20 min at RT with 10% FBS (in PBS). Subsequently, cells are again washed once with PBS.
4.
For staining, cells are incubated with the mouse anti-LAMP-2 monoclonal primary antibody (1/100 dilution in 500 $\mu$L of BSA buffer). Incubation can be performed either at RT for 1 h or at 4$^\circ$C overnight.
5.
Primary antibody solution is removed, cells are washed three times with PBS and incubated for 1 h at RT with the Alexa Fluor^® 568-conjugated secondary antibody (1/300 dilution in 500 $\mu$L of BSA buffer) (see Note 64).
6.
Finally, BSA buffer is discarded, stained cells are rinsed three times with PBS and cover slips are mounted as described above (see Subheading 3.2.2., step 7).
7.
To determine the colocalization between AV and lysosomes, confocal fluorescence microscopy should be performed at 63X magnification (see Note 65) on a device equipped with a dual FITC-Texas Red filter combination (see Notes 66 and 67).
8.
Percentage of colocalization is quantified by software-assisted analysis of green, red, and green-red merged images (see Note 68).

3.3.1.2 LysoTracker^® Probes

LysoTracker^® probes are fluorescent acidotropic, lysosomotropic, readily cell-permeant fluorochromes for labeling and tracking acidic compartments in living cells. These molecules accumulate in lysosomes and autophagosomes (as well as in other acidic subcellular compartments) due to their chemical character of weak basic amines. Since such probes are not specific for autophagosomes, they cannot be used alone to assess and/or quantify autophagy (41).

LysoTracker^® probes exist in several variants, which exhibit distinct excitation and emission spectra, to facilitate double or triple stainings. For instance, when AV are labeled with LC3-GFP (emitting in green), colocalization of AV and lysosomes can be visualized with LysoTracker^® Blue or Red.

1.
25 $\times$ 10$^{3}$ HeLa cells are cultured in 24-well plates, transfected with the LC3-GFP plasmid, and treated as described above (see Subheading 3.2.2., steps 1–4 and Notes 4– 6, 11, and 12).
2.
At the end of the desired treatments, growth medium is removed, cells are washed twice with PBS, then prewarmed medium containing the selected LysoTracker^® probe (e.g., LysoTracker^® Red) at the final concentration of 50–75 nM is added. Staining is performed at 37$^\circ$C, in 5% CO$_2$ atmosphere, for a minimal time of 30 min (see Note 69).
3.
The staining solution is removed, cells are washed once with PBS, and cover slips are mounted as described above (see Subheading 3.2.2., step 7).
4.
The cells should be observed using a confocal microscope equipped with a 63X objective (see Notes 65 and 66). A dual FITC-Texas Red filter combination is appropriate to visualize both the LC3-GFP fusion protein and LysoTracker^® Red (see Note 67).
5.
To determine the percentage of colocalization, software-assisted analysis of green, red, and red-green merged images is performed (see Note 68).

3.3.2 Mitochondria

Mitochondria are vital organelles for cellular metabolism and bioenergetics, but they are also the major regulators of cell death. Indeed, in many (if not all) paradigms of apoptosis, MMP represents the point of no return in the cascade of events that ultimately leads to the cell’s demise (42). MMP results in the leakage of potentially toxic proteins from the mitochondrial intermembrane space (IMS) into the cytosol. For instance, while cytochrome c (Cyt c) in the IMS has vital functions (by acting as an electron shuttle in oxidative phosphorylation), it participates in the activation of caspases when it is released from mitochondria (43). In addition, mitochondria play a role in stress responses and can produce reactive oxygen species (ROS) when damaged (44, 45). Selective degradation of mitochondria by autophagy is also known as “mitophagy” and is thought to be promoted by their functional impairment and/or by MMP. Mitophagy may ensure the removal of damaged and potentially dangerous mitochondria, thus acting as a quality control mechanism (46).

3.3.2.1 Transfection with pDsRed2-mito

pDsRed2-mito is a mammalian expression vector that encodes a fusion of Discosoma sp. red fluorescent protein (DsRed2)(47) and the mitochondrial targeting sequence from subunit VIII of human cytochrome c oxidase (mito)(48, 49). Upon expression, this chimeric protein readily localizes to the mitochondrial matrix, thus allowing for the visualization and tracking of mitochondria. The simultaneous detection of AV (with LC3-GFP) and mitochondria (with DsRed2-mito) provides a means to estimate the degree of mitophagy in cells.

1.
HeLa cells are seeded in 24-well plates, as described above (see Subheading 3.2.2., step 1).
2.
Twelve to 24 h after seeding, cells are co-transfected with LC3-GFP and pDsRed2-mito plasmids. For the transient transfection of cells with two plasmids, the same protocol reported above (see Subheading 3.2.2., steps 2–4, and Notes 4–6, 11, and 12) is applied. Importantly, the ratio between the total amount of plasmid DNA and Lipofectamine™ has to be maintained, so 2 $\mu$g of LC3-GFP plasmid and 2 $\mu$g of pDsRed2-mito should be employed.
3.
Following the desired treatments, cells are washed once with PBS and cover slips are mounted as detailed above (see Subheading 3.2.2., step 7).
4.
Confocal fluorescence microscopy is then performed at 63X magnification (see Notes 65 and 66). The LC3-GFP and DsRed2-mito fusion proteins can be appropriately visualized by employing a dual FITC-Texas Red filter combination (see Note 67).
5.
The degree of colocalization between green and red fluorescence is determined as described above (see Subheading 3.3.1.1. and Note 68).

3.3.2.2 MitoTracker^® Probes

MitoTracker^® probes are cell permeant, mitochondrion-selective, fluorescent stains that are concentrated by active mitochondria and well retained during fixation(50). This latter feature (which is not shared by conventional mitochondrial probes such as rhodamine 123 or tetramethylrosamine) seems to depend on the presence of a mildly thiol-reactive chloromethyl moiety. As for LysoTracker^® probes, several variants of MitoTracker^® probes exist for use in double or triple staining protocols (e.g., LC3-GFP and MitoTracker^® Red to determine the colocalization of AV and mitochondria).

1.
For colocalization studies of AV marked with LC3-GFP fusion protein and mitochondria stained with MitoTracker^® Red CMXRos, the same protocol described above for LysoTracker^® Red (see Subheading 3.3.1.2., steps 1–3, and Notes 4–6, 11, 12, and 69) is followed.
2.
In contrast to LysoTracker^® probes, MitoTracker^® probes should be employed at the final concentration of 150 nM.
3.
Confocal microscopy assessments are performed as detailed above (see Subheading 3.3.1.2.,– steps 4 and 5, and Notes 65–68).

3.3.2.3 Heat Shock 60 kDa Protein (Hsp60) Immunofluorescence Staining

In eukaryotes, Hsp60 is mainly localized to the mitochondrial matrix and is not released during cells death. Thus, it has been widely employed as a specific marker of mitochondria(51, 52).

1.
HeLa cells are cultured, transfected, treated, permeabilized, and fixed as described above for LAMP-2 staining (see Subheading 3.3.1.1., steps 1–3, and Notes 4–6, 11–16).
2.
For staining, cells are incubated with the mouse anti-Hsp60 monoclonal primary antibody (1/100 dilution in 500 $\mu$L of BSA buffer). Incubation can be performed either at RT for 1 h or at 4$^\circ$C overnight.
3.
Subsequent washing, incubation with the Alexa Fluor^® 568-conjugated secondary antibody, mounting and confocal microscopy determinations are carried out as described above (see Subheading 3.3.1.1., steps 5–8 and Notes 64–68).

3.3.3 Endoplasmic Reticulum

The ER is a network of or interconnected tubules, vesicles, and sacs involved in multiple processes, including the regulation of cytosolic calcium concentrations. Several studies suggest that the membranes of AV originate from ER. Calreticulin is the most prominent calcium-binding protein found in the ER lumen(53) and can be used as a marker in AV-ER colocalization experiments.

1.
HeLa cells are grown, transfected, treated, permeabilized, and fixed as described above for LAMP-2 staining (see Subheading 3.3.1.1., steps 1–3 and Notes 4–6, 11–16).
2.
For staining, cells are incubated with the rabbit anti-calreticulin polyclonal primary antibody (1/100 dilution in 500 $\mu$L of BSA buffer). Incubation can be performed either at RT for 1 h or at 4$^\circ$C overnight.
3.
Subsequent washing, staining with the Alexa Fluor^® 568-conjugated secondary antibody, mounting, and confocal microscopy determinations are carried out as described above (see Subheading 3.3.1.1., steps 5–8 and Notes 64–68).

3.4 Methods for Measuring Cell Death–Related Parameters

Most cell death in vertebrates proceeds via the intrinsic, mitochondrial pathway of apoptosis (54). In several models of cell death, MMP is considered as the point of no return in the pro-apoptotic cascade of events (42). MMP may result from the activity of pro-apoptotic members of the Bcl-2 family (such as Bax, Bak, and BH3-only proteins like Bid)(55, 56) or from the opening of a multiprotein complex, the permeability transition pore complex (PTPC)(57, 58). The sudden increase in permeability of the mitochondrial inner membrane (IM) that derives from PTPC opening (a process known as mitochondrial permeability transition, MPT) leads to dissipation of the mitochondrial transmembrane potential$\Delta\Psi_{\rm m}$ and eventually to the rupture of the mitochondrial outer membrane (OM) (42). Irrespective of its initiation (be it mediated by Bcl-2 pro-apoptotic proteins or by opening of the PTPC), MMP leads to the functional impairment of mitochondria and to the release of IMS proteins into the cytosol. These include caspase activators like Cyt c(43, 59), Omi/HtrA2 (Omi stress-regulated endoprotease/high temperature requirement protein A 2)(60) and Smac/DIABLO (second mitochondria-derived activator of caspase/direct IAP binding protein with a low pI)(61, 62), as well as caspase-independent cell death effectors such as apoptosis-inducing factor (AIF)(63, 64) and endonuclease G (EndoG)(65).

The assessment of early mitochondrial alterations allows for to identification of cells that are committed to death but have not yet displayed an apoptotic phenotype. The $\Delta\Psi_{\rm m}$ (providing direct indications on the IM permeability status) can be assessed by means of cationic fluorescent probes (66). Such lipophilic dyes (e.g., DiOC$_6$(3), TMRM) accumulate in mitochondria driven by $\Delta\Psi_{\rm m}$, according to the Nernst equation, and can be used in situ for microscopic assessments of $\Delta\Psi_{\rm m}$ as well as in cytofluorometric-based procedures for $\Delta\Psi_{\rm m}$ quantification (52).

OM permeabilization can be assessed by the detection of IMS proteins (e.g., Cyt c, AIF) in the cytosol. This can be achieved by immunoblotting subcellular fractions with antibodies against Cyt c or AIF, or by in situ immunofluorescence microscopy on fixed and permeabilized cells (51, 67, 68). As an alternative, cells can be transfected with cDNA constructs encoding IMS proteins fused to a GFP moiety(69, 70). Such chimeric proteins are targeted to IMS as their normal counterparts. Upon apoptosis stimulation, videomicroscopy permits to follow the redistribution of GFP-tagged proteins from IMS to other subcellular compartments in real time. Among these techniques, immunofluorescence microscopy seems the most appropriate to detect the release of IMS protein, since it provides more accurate information on their redistribution than subcellular fractionation followed by immunoblotting, and it is not associated with the technical requirements of videomicroscopy(52).

One of the classical hallmarks of apoptosis is the externalization of phosphatidylserine (PS) and phosphatidylethanolamine (PE)(71, 72). In normal conditions, these phospholipids are sequestered on the cytoplasmic leaflet of the plasma membrane, but they translocate to the outer leaflet upon apoptosis induction. Fluorescent Annexin V that specifically binds to PS exposed on the extracytoplasmic side of the plasma membrane can be used to identify apoptotic cells.

Following MMP, mitochondrial uncoupling results in enhanced generation of ROS. This alteration may be quantified (in association with other mitochondrial parameters like the $\Delta\Psi_{\rm m}$) by means of cytofluorometric techniques. For instance, the nonfluorescent probe hydroethidine (HE) is oxidized by ROS to ethidium bromide (EthBr), which emits red fluorescence(73). In particular, HE is more sensitive to superoxide anion than 2$^\prime$,7$^\prime$-dichlorofluorescein diacetate, which preferentially measures H$_{2}{\rm O}_{2}$ formation (45, 73, 74). Alternatively, ROS-induced damage of mitochondria can be determined indirectly, by assessing the oxidation state of cardiolipin, a lipid restricted to the IM. This may be achieved with nonyl acridine orange (NAO), a fluorochrome that specifically interacts with nonoxidized, intact cardiolipin (75). Consequently, a reduction in NAO fluorescence is an indicator of ROS-mediated cardiolipin oxidation.

Downstream MMP, the activation of the caspase cascade is frequently associated with the onset of apoptotic cell death(42, 54). However, caspase-independent routes to death have been reported in several models of apoptosis(76). Moreover, caspase activation occurs in several nonapoptotic scenarios (77). As it stands, the detection of activated caspases cannot be used alone to assess apoptosis, but it may provide a useful complementary assay to other techniques. Among the various caspases that execute the apoptotic program, a prominent one is caspase-3 (78). Hence, detection of the large fragment (17/19 kDa) of active caspase-3 (caspase-3a), resulting from the caspase-9–mediated cleavage adjacent to Asp175, is commonly considered as an indicator of apoptosis induction.

The integrity of plasma membrane represents a central difference between apoptosis and necrosis. During primary necrosis, the generalized swelling of the cell and of cytoplasmic organelles results in the early breakdown of plasma membrane. On the contrary, plasma membrane integrity is maintained until the late stages of apoptosis, before secondary necrosis occurs (22). Thus, the vital dye propidium iodide (PI, emitting in red), which cannot enter living cells nor cells undergoing apoptosis, can be used to differentiate between apoptotic and necrotic cells by means of flow cytometry. Also, other dyes that bind stoichiometrically to double-stranded DNA, such as 4$^\prime$,6-diamidino-2-phenylindole dihydrochloride (DAPI) and Hoechst 33342, can be used to measure cell viability. Moreover, these molecules can be employed to label the nuclei of permeabilized cells for fluorescence-based assessments. When examined by fluorescence microscopy, nuclei from normal cells glow brightly and homogeneously and can be easily differentiated from apoptotic nuclei, which display chromatin condensation (nuclear pyknosis, at early stages) or a total segmented morphology (karyorhexis, later during the process(22).

3.4.1 Analysis of $\Delta\psi_{m}$ Dissipation

1.
25 $\times$ 10$^{3}$ HeLa cells are seeded in 24-well plates (0.5 mL of growth medium per well). After 12–24 h, cells are treated with the desired stimuli.
2.
At the end of treatments, cells are collected by trypsinization (300 $\mu$L of trypsin/EDTA per well), and spun at 300g for 5 min (RT) (see Note 70).
3.
Supernatant is discarded and cells are labeled at 37$^\circ$C either with 150 nM TMRM (emitting in red) or with 40 nM DiOC$_6$(3) (emitting in green) in 200 $\mu$L of growth medium (see Notes 71 and 72).
4.
Cytofluorometric acquisitions are performed by means of a conventional cytofluorometer equipped with a single laser for excitation (see Notes 73 and 74).
5.
Analysis should be performed on viable, normal-sized cells by gating a single population with normal forward and side scatter parameters (see Note 75). Each experiment should be supported by an appropriate set of controls (see Note 76).

3.4.2 Analysis of Phosphatidylserine Externalization

Apoptosis-related PS translocation to the outer leaflet of plasma membrane is detected by staining the cells with fluorescein isothiocyanate (FITC)-conjugated Annexin V (see Note 77).

1.
HeLa cells are cultured, treated, and collected as described above (see Subheading 3.4.1., steps 1 and 2, and Note 70).
2.
Supernatant is discarded and cells are washed in 1 mL of binding buffer and centrifuged at 300g for 10 min (RT). Supernatant is removed completely and the washing step is repeated.
3.
Cells are resuspended in 100 $\mu$L of binding buffer and 10 $\mu$L of Annexin V-FITC are added.
4.
Samples are carefully mixed and incubated at RT under protection from light for 15 min (see Notes 78 and 79).
5.
Cells are washed by adding 1 mL binding buffer and spun at 300 g for 10 min (RT). Supernatant is discarded and cells are resuspended in 500 $\mu$L of binding buffer (see Note 80).
6.
Immediately following, cytofluorometric acquisitions are performed as described above (see Subheading 3.4.1., steps 4 and 5, and Notes 72–73, 75, 80, and 81).

3.4.3 Determination of Mitochondrial ROS Generation and Local ROS Effects

1.
100 $\times$ 10$^{3}$ HeLa cells are cultured in 12-well plates (1 mL of growth medium).
2.
Twelve to 24 h later, the desired treatments are administered to the cultures.
3.
At the end of stimulation, cells are trypsinized (500 $\mu$L trypsin/EDTA per well), and spun at 300g for 5 min (RT) (see Note 70).
4.
Upon removal of the supernatant, cells are labeled either with 25 $\mu$ M HE or with 100 nM NAO in 500 $\mu$L of culture medium.
5.
Labeling is performed at 37$^\circ$C for 10 min.
6.
Then, cytofluorometric acquisitions are performed as described above (see Subheading 3.4.1., steps 4 and 5, and Notes 72, 73, 75, 82, and 83).

3.4.4 Determination of the Mitochondrial Release of IMS Proteins by Immunofluorescence Microscopy

1.
25 $\times$ 10$^{3}$ HeLa cells are seeded in 24-well plates as described above (see Subheading 3.1.1., step 1). After 12–24 h, cells are treated with the desired stimuli.
2.
Following treatments, cells are fixed and permeabilized as described before (see Subheading 3.3.1.1., steps 2 and 3 and Notes 13–16).
3.
For double staining, cells are then incubated with the rabbit anti-AIF and with the mouse anti-cytochrome c primary antibodies (both at 1/100 dilution in 500 $\mu$l of BSA buffer). Staining can be performed indifferently at RT for 1 h or at 4$^\circ$C overnight (see Note 84).
4.
Subsequent washing, staining with anti-mouse and anti-rabbit Alexa Fluor^®-conjugated secondary antibodies, and mounting are performed as described above (see Subheading 3.3.1.1., steps 5 and 6 and Note 64).
5.
Examination is then carried out at the fluorescence microscope by using an oil immersion objective (50–65$\times$ magnification). Excitation and emission filters should be selected according to the following absorption/emission peaks: 495/519 nm for Alexa Fluor^® 488 (green emission); 578/603 nm for Alexa Fluor^® 568 (red emission); 352/461 nm for Hoechst 33342 (blue emission).

A distinct, punctuate cytoplasmic pattern of fluorescence indicates the mitochondrial localization of AIF (red fluorescence) and Cyt c (green fluorescence). This can be verified by counterstaining mitochondria with antibodies directed against sessile mitochondrial markers, including OM integral (e.g., voltage-dependent anion channel) or matrix proteins (e.g., Hsp60). As an alternative, mitochondria can be prestained with MitoTracker^® probes, as detailed above (see Subheading 3.3.2.2.).

On the contrary, a diffuse staining pattern can be observed upon the mitochondrial release of IMS proteins. Following the release from mitochondria, Cyt c is found mainly in the cytosol (43), whereas AIF accumulates in the nucleus (63, 79). The percentage of cells exhibiting a redistribution of AIF and Cyt c should be determined in a statistically meaningful population (see Note 19).

3.4.5 Caspase 3$_{a}$ Staining

1.
HeLa cells are seeded in 24-well plates, treated, permeabilized, and fixed as detailed above (see Subheading 3.1.1., step 1, Subheading 3.3.1.1., steps 2 and 3, and Notes 13–16).
2.
For staining, cells are then incubated with the rabbit anti-caspase-3a (1/100 in 500 $\mu$L of BSA buffer). Staining can be performed indifferently at RT for 1 h or at 4$^\circ$C overnight (see Note 84).
3.
Subsequent washing, staining with anti-rabbit Alexa Fluor^®-conjugated secondary antibodies, mounting, and examination are performed as previously detailed (see Subheading 3.3.1.1., steps 5 and 6 and Note 64, Subheading 3.4.4., step 5).
4.
Caspase-3 activation results in a bright, diffuse cytoplasmic red fluorescence, markedly in contrast with the background signal of negative cells. For each sample, the percentage of positivity for caspase-3a should be quantified in a cell population of statistical relevance (see Note 19).

3.4.6 Detection of Plasma Membrane Integrity

Dead cells incorporate the so-called vital dyes, that are normally excluded by the barrier given by an intact plasma membrane. One of the most widely used vital dyes is propidium iodide (PI).

1.
HeLa cells are cultured, treated, and collected as described above (see Subheading 3.4.1., steps 1 and 2 and Note 70).
2.
Supernatant is discarded and cells are labeled at 37$^\circ$C with 1 $\mu$g/mL PI in 200 $\mu$L of growth medium (see Note 71).
3.
Then, cytofluorometric acquisitions are performed as described above (see Subheading 3.4.1., steps 3–5 and Notes 72, 73, 75, 80, and 85).

3.5 A General Strategy for the Resolution of the Central Silemma of Autophagic Cell Death

As outlined in the introduction to this chapter, one of the major problems raised by the observation of autophagic cell death (type 2 cell death) concerns causality. Does cell death simply occur with autophagy, which would constitute a late and desperate attempt of the cell to adapt to stress? Or is cell death mediated through autophagy, meaning that self-eating is one of steps that leads to self-killing? In the case of cells presenting massive autophagic vacuolization followed by plasma membrane barrier and disintegration, cell death should be considered as occurring through autophagy sensu stricto if all the following requirements are met:

1.
Cell death (loss of membrane integrity with positive staining for PI or similar vital dyes) should be preceded by all signs of autophagy (double-membraned vesicles, LC3-GFP aggregation, LC3 proteolytic maturation, increase of the acidic subcellular compartment, enhanced turnover of long-lived proteins, co-localization of lysosomal and mitochondrial or ER markers), and these signs should be inhibitable by siRNAs targeting essential proteins of the process (atg5, atg6/Beclin-1, atg10, atg12, hVps34).
2.
Cell death should occur without unequivocal morphological manifestations of apoptosis (e.g., pyknosis, karyorhexis, shrinkage, and formation of the apoptotic bodies). However, cell death may be linked at late stages to mitochondrial permeabilization and caspase activation. These alterations would indicate that apoptotic effector mechanisms have been activated.
3.
Most importantly, cell death in all its possible manifestations (autophagic, apoptotic, or necrotic) should be inhibited by genetic manipulations designed to reduce or suppress the expression of atg genes or hVps34. Thus, the inhibition of autophagy by specific methods (as opposed to the unspecific pharmacological modulators that are currently available) should prevent cell death and maintain the cells healthy and viable.

If it is possible to substantiate by means of these methods that cells can effectively die through autophagy, then it will become a challenging endeavor for future investigation to determine the functional importance of autophagic cell death in tissue homeostasis, remodeling, and pathology.

4 Notes

1.
Working dilutions of the inducers and inhibitors of autophagy should be prepared from the stock solutions shortly before use. Intermediate dilutions, when necessary, can be made either in sterile PBS or in growth medium.
2.
The minimal time necessary to observe a stimulation of the autophagic process varies according to the inducer from 1 h (e.g., upon stimulation with xestospongin C) up to 12 h (e.g., after depletion of serum and amino acids). Thus, to avoid excessive toxicity, it is recommendable NOT to incubate cells for longer than 36–48 h prior to the quantification of autophagic vacuoles.
3.
Contrarily to the manufacturer’s instructions (recommending 30–50% of confluence), we found that optimal transfection rates are attained when cells are slightly more confluent (50–70%). However, excessive confluence (at levels coinciding with a reduction of proliferation) should be carefully avoided, since it results in a significant drop of the efficacy of transfection.
4.
When the diluted Oligofectamine™ and the siRNA solution are mixed, the solution may appear cloudy. The same applies to Lipofectamine™ and plasmid solutions.
5.
The liposome-mediated siRNA transfection protocol is carried out entirely at RT under a common safety cabinet. However, it is recommendable to keep the tubes containing the stock solutions of siRNAs and Oligofectamine™ in an ice bath (and to return them to storage conditions immediately after use), to avoid the degradation of reagents, and to minimize solvent evaporation (both of which may eventually affect the concentration of the stocks). The same applies to the liposome-mediated transfection of plasmids.
6.
The absence of serum is especially important for optimal formation of the Oligofectamine™:siRNA complexes. Nevertheless, contrary to common beliefs, we found that complexes (once properly assembled in serum-free conditions) can be added to cells cultured in complete growth medium without relevant reductions of the transfection efficiency. This practice removes the need for serum addition 4 h after transfection. The same applies to the liposome-mediated transfection of plasmids.
7.
Given the very high affinity of transfection complexes for the plasma membrane, they should be added to cells dropwise, by trying to cover the whole surface of the growth medium, in order to avoid intrawell variations of the transfection efficiency.
8.
Due to multiple variables, including the efficacy of different siRNAs (also targeting the same transcript) as well as the half-life of target proteins, the time required for optimal knock-down may range from a few hours to several days. As a guideline, we recorded satisfactory levels of downregulation of several proteins involved in autophagy (e.g., Beclin-1, LAMP-2, Atg5, Atg10, Atg12, hVps34) 48 h after transfection. However, it is recommendable to check the kinetics of protein downregulation for each siRNA and specific experimental setting. This should be accomplished both at the mRNA level (by RT-PCR, using the appropriate primers) and at the protein level (by immunoblotting with the specific antibodies).
9.
As a negative control, cells transfected with scrambled siRNAs or with siRNAs targeting irrelevant sequences (not found in the cellular genome) should be carried along the experimental procedure and subjected to the same treatments administered to cells transfected with siRNA targeting proteins essential for autophagy.
10.
At this stage, the seeding concentration heavily depends on the duration and strength of the subsequent treatments. As a guideline, for treatments of 24 h or less (administered 24 h after plating) we use seed 100 $\times$ 10$^{3 }$cells in 1 mL of growth medium (12-well plates) or 25 $\times$ 10$^{3}$ cells in 0.5 mL of growth medium (24-well plates).
11.
We recorded the expression of the LC3-GFP fusion protein already 12 h after transfection. Faster expression is not excluded, but should be controlled in the specific experimental setting.
12.
As for siRNAs, an adequate control has to be carried along the entire experimental procedure and treated as samples (cells transfected with the plasmid encoding the LC3-GFP chimeric protein). To this aim, untransfected cells may be used. However, a more stringent control is provided by cells transfected with the empty cloning vector.
13.
PFA is toxic by inhalation and should be handled under an appropriate fume hood.
14.
Stability of the PFA solution is limited. Accordingly, it should be prepared shortly before use. Once dissolved, it is recommendable to keep PFA on ice bath throughout the experiment.
15.
Picric acid (chemical name 2,4,6-trinitrophenol) is a close derivative of trinitrotoluene (TNT) and is toxic and explosive if allowed to dry out. Thus, maximal care should be used when handling picric acid.
16.
At washing steps, gentle pipetting is recommended to avoid the detachment of cells from the coverslips.
17.
Hoechst 33342 is carcinogenic and mutagenic.
18.
While wild-type GFP has two excitation peaks, a major one at 395 nm (in the long UV range) and a smaller one at 478 nm (blue), the red-shifted mutant variant encoded by the plasmid pEGFP-C1 (cloning vector from Clontech Laboratories, Palo Alto, CA) is characterized by a single absorption peak at 488 nm and higher expression in mammalian cells.
19.
For quantification purposes, the frequency of cells exhibiting cytoplasmic LC3-GFP aggregation should be determined among (at least) 200 cells. Moreover, the number of AV (green dots) per cell should be counted in a representative sample of (at least) 50 cells. The same considerations about the population size apply to other techniques for AV quantification (e.g., MDC or CMFDA staining), as well as to the assessment of translocation of IMS proteins and caspase-3 activation.
20.
L-[U-$^{14}$C]-Valine is a radiochemical. Its handling and storage should conform to the current safety rules.
21.
The prechase period is crucial to get rid of short-lived proteins, which are indeed degraded during this step.
22.
At this stage, the use of an autophagy inhibitor provides the required negative control.
23.
MDC is an autofluorescent molecule characterized by a relatively weak emission, peaking at 525 nm. However, it has been demonstrated that it behaves as a solvent polarity probe (80) and that its interaction with membrane lipids (as occurring in AV) enhances by severalfold its emission, while shifting its emission to 498 nm.
24.
To distinguish the characteristic vesicular distribution of MDC in autophagosomes, each experiment should be performed and scored with the appropriate controls. To this aim, NF medium combined or not with an autophagy inhibitor of choice may provide positive and negative control conditions, respectively.
25.
CMFDA is characterized by an absorption and fluorescence emission maxima of 492 and 517 nm, respectively, as determined in aqueous buffer or methanol. Values may exhibit slight variations in cellular environments.
26.
To minimize proteolytic degradation, after washing samples should be kept in ice bath until the addition of loading buffer and boiling or storage.
27.
Lysis should be performed in the minimal volume of lysis buffer that allows for total resuspension of the pellets (usually 30 $\mu$L will suffice). Lysis may be facilitated by thorough pipetting or quick vortexing. Keeping the lysis buffer volume to a minimum reduces the dilution of the protein content of samples, thus resulting in easier gel loading (see also Note 31).
28.
Lysates may be stored at –20$^\circ$C for prolonged periods (several months) without significant degradation of protein content.
29.
Protein quantification is performed by interpolating a calibration curve that is built from a serial dilution of BSA. BSA standards are rarely (if ever) included in the kits for quantification and should be prepared shortly before use from the BSA stock solution.
30.
The amount of proteins that should be loaded onto the polyacrylamide gel for separation depends on the size and expected expression levels of protein of interest. For large or highly expressed proteins, 20 $\mu$g may suffice to visualize the corresponding band upon immunoblotting. For small or rare proteins larger amounts (up to 60–80 $\mu$g) are required.
31.
When the quantity of proteins for loading has been determined, samples that will be separated on the same gel are prepared in the minimal volume, which allows one to load the same amount of proteins for each lysate and the addition of 5X loading buffer (to a final concentration of 1X). Keeping the loading volume to a minimum facilitates the loading procedure. In case of diluted samples, 6–7X loading buffer may be employed.
32.
Boiling is performed to complement the denaturating activity of lysis and loading buffers (containing SDS, reducing agents, and calcium chelators), with the aim to break the inter- and intramolecular bonds responsible of the higher-level (tertiary and quaternary) protein structures. This step ensures that the subsequent separation is truly based on the size of protein subunits, with little influence of protein–protein interaction and native structural features.
33.
The percentage of acrylamide directly determines the sieving properties of the gel, thus influencing its separation range and resolution. As a guideline, proteins with a molecular weight higher than 100 kDa should be resolved on 5–10% total acrylamide gels, whereas smaller proteins can be separated on gels containing 10–15% acrylamide. To facilitate pouring and subsequent handling, concentrations of acrylamide higher than 15% should be avoided.
34.
Monomeric acrylamide is a neurotoxin and suspected carcinogen. Avoid skin contact and inhalation. Always wear gloves and protective clothing and handle nonpolymerized acrylamide solutions under a fume hood.
35.
In polyacrylamide gels, TEMED acts as crosslinking agent. Also its concentration influences the final properties of the polymerized gel.
36.
TEMED is toxic by inhalation. It should be always handled under a fume hood.
37.
APS stock solution (10% in dH$_2$O) is unstable. Storage at 4$^\circ$C is possible, but never exceed one week. The use of old APS solutions may result in incomplete polymerization of the acrylamide gel.
38.
APS generates the free radicals required for the polymerization of acrylamide, thus acting as the catalyst of the reaction. Accordingly, APS should be added as the last component of the gel solution.
39.
This protocol reports the amounts of reagents required for a single gel. These quantities may be readily scaled up to pour more gels from the same starting solution. However, no more than four to six gels should be cast from the same solution, to avoid premature polymerization.
40.
Hint: Upon pouring, the gel solution remaining in the tube should not be immediately discarded, but left in contact with air. It will help in determining the polymerization state of the gels.
41.
Pouring should be performed quickly, while avoiding the formation of bubbles that would interfere with protein migration.
42.
It is recommended to leave at least the upper third of the glass height for the stacking gel.
43.
Isopropanol may be substituted with pure water.
44.
Polymerization time depends on acrylamide concentration, but usually 10–15 min are sufficient. Prolonged polymerization time or incomplete polymerization may result from an old APS stock (see also Note 38). Polymerization can be verified by examining the remnant solution in the tube in which the gel solution has been prepared (see also Note 40).
45.
The use of a stacking gel improves the resolution of electrophoresis because it causes proteins to accumulate and stack as a very thin zone at the stacking gel/running gel boundary and because it arranges the proteins according to their order of mobility before their entry into the running gel.
46.
The comb (which determines the size of wells) should be selected according to the number and final volume of the samples. 5-, 9-, 10-, and 15-well combs are available, allowing to load approximately 60, 50, 40, and 20 $\mu$L of samples, respectively.
47.
Importantly, the migration buffer should completely cover the wells but not create a communication between the upper and lower chambers that would compromise the correct electrical flow in the cell.
48.
Prestained markers covering different ranges of MW are commercially available and allow for visual follow-up of migration.
49.
Before entry in the running gel, voltage should be set at 50–70 mV. Thereafter, voltage can be augmented to 140 mV. It should be kept in mind that higher voltages result in reduced migration time but also in lower resolution.
50.
Migration fronts, as indicated by the fast migrating bromophenol blue component of the loading buffer, may be safely run off the gel, if desired.
51.
During assembly, special care should be taken to avoid the formation of bubbles (which will interfere with the transfer of proteins) between the various layers of the “sandwich.” To remove bubbles, a common 5-mL pipet can be gently rolled on top of each component, before the addition of the next one.
52.
Each transfer cassette allows the insertion of two “sandwiches.” To avoid the loss of proteins during the transfer, they should both be oriented in the same fashion (see also Note 53).
53.
If the cassette is inserted with the reverse orientation (membrane between the gel and the cathode), negatively charged proteins will be attracted by the anode across the filter paper sheets and the sponge and eventually be lost in the transfer buffer.
54.
Due to the intense current, the transfer apparatus temperature tends to increase excessively, if not controlled. As an alternative to the ice block, the entire transfer procedure can be performed in a cold room (4$^\circ$C).
55.
As an alternative, the transfer can be accomplished overnight at a constant current of 45 mA per membrane.
56.
To keep the buffer temperature as homogeneous as possible, the transfer should be performed on top of a magnetic stirrer (gentle stirring).
57.
Upon careful rinsing in PBS or dH$_2$O, sponges can be re-used. Filter papers are discarded.
58.
At this stage, problems during the transfer (such as bubbles) can be easily identified. Also, membranes can be wrapped in transparent plastic film and photocopied (or digitalized by scanning) to keep track of the correct transfer.
59.
At this step, attention should be paid to ensure the even coverage of the membrane with the chemiluminescent substrate.
60.
Photographic films are light sensitive and should be handled only in the dark room until the end of the development processing.
61.
Exposure time varies greatly according to the intensity of signals from the HRP-catalyzed chemiluminescent reaction. In turn, this depends on several factors, including the amount of protein in the band, the affinity of primary and secondary antibodies, and the time passed from the incubation of the chemiluminescent substrate. For most proteins, exposure times between 1 and 5 min provide good results. When proteins are small or scarcely expressed (or when the antibodies display reduced affinity), longer exposures (up to 10–12 hours) should be attempted. On the contrary, when proteins are large or well represented in the sample (or when the antibodies exhibit high affinity), a few seconds may be sufficient. Some trials may be required to obtain the correct exposition time. Given the time-dependent decay of the luminescent signal, short exposures should be performed before longer ones.
62.
If the band of interest cannot be detected upon overnight exposure, the reagent Supersignal West Femto Maximum Sensitivity Substrate may provide enhanced sensitivity.
63.
Loading control should be performed with an antibody directed against a highly represented protein, whose amount is stable in most experimental settings. Usually, GAPDH, $\alpha$-tubulin, or $\beta$-actin are detected as loading control.
64.
To avoid photobleaching of the fluorescent antibodies, samples should be protected from light from this step onward.
65.
As an alternative, 40 or 100X objectives may also be employed.
66.
We detected the colocalization of two fluorescent signals by means of a Zeiss LSM 510 confocal microscope.
67.
The dual FITC-Texas Red filter combination allows for the visualization of LC3-GFP (absorption/emission peaks at 488/507 nm) and DsRed2-mito (558/583 nm) fusion proteins, as well as of LysoTracker^® Red (577/590 nm), MitoTracker^® Red CMXRos (579/599 nm), and the Alexa Fluor^® 488 (495/517) and 568 (578/603 nm) fluorochromes.
68.
To this aim, we routinely use the free, Java^®-based, open-source software ImageJ.
69.
Staining times may be extended up to 2 h, according to cell type and/or general conditions of the culture.
70.
At this stage, the supernatant of treated cells should not be discarded, to avoid the loss of part of the population detached from the plate upon the treatment. Rather, supernatants should be collected in FACS tubes, to which the corresponding cell suspension will be added after trypsinization.
71.
To avoid probe-dependent toxicity to the cells, cytofluorometric acquisitions should be performed within 30 min.
72.
When large series of samples are to be analyzed (>12 tubes), the interval between labeling and cytofluorometric analysis should be kept constant.
73.
To this aim, we routinely use a Becton Dickinson FACScan cytofluorometer, equipped with an argon ion laser emitting at 488 nm. The following channels are employed for the detection of fluorescent emissions: FL1 for DiOC$_6$(3), FITC and NAO (also in FL2); FL2 for NAO (also in FL1) and TMRM; FL3 for PI and EthBr (as resulting from HE superoxide anion-mediated oxidation) (see also Notes 74, 80, and 81).
74.
DiOC$_6$(3) is characterized by absorption/emission peaks at 484/501 nm, TMRM by maxima at 543/573 nm.
75.
As a reminder, the forward light scatter gives an indication on cell size whereas the side scatter is related to several factors, including granularity, refractive index, and shape of the cells.
76.
Since the incorporation of DiOC$_6$(3) and TMRM can be influenced by parameters not related to mitochondria (e.g., cell size, plasma membrane permeability, activity of multidrug resistance pumps, etc.) (45, 66), results can only be interpreted when the difference in staining profiles between different experimental conditions (e.g., control vs. apoptosis) are NOT affected by the incubation with a ionophore causing the complete disruption of $\Delta\Psi_{\rm m}$. Hence, an adequate set of controls can be prepared by splitting in 2 aliquots each sample, and by preincubating one of the two series with 50–100 $\mu$M CCCP for 10–15 min before staining (37$^\circ$C, 5% CO$_2$).
77.
To this aim, we routinely employ the Annexin V-FITC kit from Miltenyi Biotec, according to the manufacturer’s instructions.
78.
Protection from light prevents photobleaching of FITC fluorochrome.
79.
Higher temperatures or longer incubation times lead to nonspecific labeling.
80.
If desired, immediately before cytofluorometric acquisition, 5 $\mu$L of a 100 $\mu$g/mL solution of PI can be added for the simultaneous detection of cells with ruptured plasma membranes. PI is characterized by absorption/emission peaks at 535/617 nm (see also Note 85).
81.
FITC exhibits an absorption maximum at 494 nm and an emission one at 519 nm.
82.
EthBr resulting from the ROS-mediated conversion of HE readily binds to DNA, and this complex is characterized by absorption/emission peaks of 300/603 nm (as opposite to reduced HE, emitting at 420 nm).
83.
NAO displays absorption/emission peaks at 500/520 nm.
84.
At the same time, nuclear counterstaining can be performed by the addition of 2 $ \mu $ M Hoechst 33342, together with primary antibodies.
85.
PI staining may be associated with either DiOC$_6$(3) or Annexin V to distinguish normal cells (PI$^{-}$, DiOC$_6$(3)$^{\rm high}$, or Annexin V$^{-}$), dying cells (PI$^{-}$, DiOC$_6$(3)$^{\rm low}$, or Annexin V$^{+}$), and dead cells (PI$^{+}$) within the same population.

References

Klionsky, D. J., and Emr, S. D. (2000) Autophagy as a regulated pathway of cellular degradation. Science 290, 1717–1721.
Article CAS PubMed Google Scholar
Mizushima, N., Ohsumi, Y., and Yoshimori, T. (2002) Autophagosome formation in mammalian cells. Cell Struct. Funct. 27, 421–429.
Article PubMed Google Scholar
Boya, P., Gonzalez-Polo, R. A., Casares, N., et al. (2005) Inhibition of macroautophagy triggers apoptosis. Mol. Cell Biol. 25, 1025–1040.
Article CAS PubMed Google Scholar
Klionsky, D. J. (2005) The molecular machinery of autophagy: unanswered questions. J. Cell Sci. 118, 7–18.
Article CAS PubMed Google Scholar
Ohsumi, Y. (2001) Molecular dissection of autophagy: two ubiquitin-like systems. Nat Rev Mol. Cell Biol. 2, 211–216.
Article CAS PubMed Google Scholar
Kabeya, Y., Mizushima, N., Ueno, T., et al. (2000) LC3, a mammalian homologue of yeast Apg8p, is localized in autophagosome membranes after processing. EMBO J. 19, 5720–5728.
Article CAS PubMed Google Scholar
Kihara, A., Kabeya, Y., Ohsumi, Y., and Yoshimori, T. (2001) Beclin-phosphatidylinositol 3-kinase complex functions at the trans-Golgi network. EMBO Rep. 2, 330–335.
Article CAS PubMed Google Scholar
Pattingre, S., Tassa, A., Qu, X., et al. (2005) Bcl-2 antiapoptotic proteins inhibit Beclin 1-dependent autophagy. Cell 122, 927–939.
Article CAS PubMed Google Scholar
Mizushima, N. (2005) The pleiotropic role of autophagy: from protein metabolism to bactericide. Cell Death Differ. 12 (Suppl. 2), 1535–1541.
Google Scholar
Yorimitsu, T., and Klionsky, D. J. (2005) Autophagy: molecular machinery for self-eating. Cell Death Differ. 12 (Suppl. 2), 1542–1552.
Google Scholar
Golstein, P., and Kroemer, G. (2007) Cell death by necrosis: towards a molecular definition. Trends Biochem. Sci., in press.
Google Scholar
Gonzalez-Polo, R. A., Boya, P., Pauleau, A. L., et al. (2005) The apoptosis/autophagy paradox: autophagic vacuolization before apoptotic death. J. Cell. Sci. 118, 3091–3102.
Article CAS PubMed Google Scholar
Yamamoto, A., Tagawa, Y., Yoshimori, T., Moriyama, Y., Masaki, R., and Tashiro, Y. (1998) Bafilomycin A1 prevents maturation of autophagic vacuoles by inhibiting fusion between autophagosomes and lysosomes in rat hepatoma cell line, H-4-II-E cells. Cell Struct. Funct. 23, 33–42.
Article CAS PubMed Google Scholar
Lum, J. J., Bauer, D. E., Kong, M., et al. (2005) Growth factor regulation of autophagy and cell survival in the absence of apoptosis. Cell 120, 237–248.
Article CAS PubMed Google Scholar
Boya, P., Gonzalez-Polo, R. A., Poncet, D., et al. (2003) Mitochondrial membrane permeabilization is a critical step of lysosome-initiated apoptosis induced by hydroxychloroquine. Oncogene 22, 3927–3936.
Article CAS PubMed Google Scholar
Yu, L. Y., Jokitalo, E., Sun, Y. F., et al. (2003) GDNF-deprived sympathetic neurons die via a novel nonmitochondrial pathway. J. Cell Biol. 163, 987–997.
Article CAS PubMed Google Scholar
Mills, K. R., Reginato, M., Debnath, J., Queenan, B., and Brugge, J. S. (2004) Tumor necrosis factor-related apoptosis-inducing ligand (TRAIL) is required for induction of autophagy during lumen formation in vitro. Proc. Natl. Acad. Sci. USA 101, 3438–3443.
Article CAS PubMed Google Scholar
Yu, L., Alva, A., Su, H., et al. (2004) Regulation of an ATG7-beclin 1 program of autophagic cell death by caspase-8. Science 304, 1500–1502.
Article CAS PubMed Google Scholar
Shimizu, S., Kanaseki, T., Mizushima, N., et al. (2004) Role of Bcl-2 family proteins in a non-apoptotic programmed cell death dependent on autophagy genes. Nat. Cell Biol. 6, 1221–1228.
Article CAS PubMed Google Scholar
Pyo, J. O., Jang, M. H., Kwon, Y. K., et al. (2005) Essential roles of Atg5 and FADD in autophagic cell death: dissection of autophagic cell death into vacuole formation and cell death. J. Biol. Chem. 280, 20722–20729.
Article CAS PubMed Google Scholar
Marchetti, P., Castedo, M., Susin, S. A., et al. (1996) Mitochondrial permeability transition is a central coordinating event of apoptosis. J. Exp. Med. 184, 1155–1160.
Article CAS PubMed Google Scholar
Kroemer, G., El-Deiry, W. S., Golstein, P., et al. (2005) Classification of cell death: recommendations of the Nomenclature Committee on Cell Death. Cell Death Differ. 12 (Suppl. 2), 1463–1467.
Google Scholar
Debnath, J., Baehrecke, E. H., and Kroemer, G. (2005) Does autophagy contribute to cell death? Autophagy 1, 66–74.
Google Scholar
Codogno, P., and Meijer, A. J. (2005) Autophagy and signaling: their role in cell survival and cell death. Cell Death Differ. 12 (Suppl. 2), 1509–1518.
Google Scholar
Lum, J. J., DeBerardinis, R. J., and Thompson, C. B. (2005) Autophagy in metazoans: cell survival in the land of plenty. Nat. Rev. Mol. Cell Biol. 6, 439–448.
Article CAS PubMed Google Scholar
Sarbassov, D. D., Ali, S. M., and Sabatini, D. M. (2005) Growing roles for the mTOR pathway. Curr. Opin. Cell Biol. 17, 596–603.
Article CAS PubMed Google Scholar
Castedo, M., Ferri, K. F., and Kroemer, G. (2002) Mammalian target of rapamycin (mTOR): pro- and anti-apoptotic. Cell Death Differ. 9, 99–100.
Article CAS PubMed Google Scholar
Asnaghi, L., Bruno, P., Priulla, M., and Nicolin, A. (2004) mTOR: a protein kinase switching between life and death. Pharmacol. Res. 50, 545–549.
Article CAS PubMed Google Scholar
Petiot, A., Ogier-Denis, E., Blommaart, E. F., Meijer, A. J., and Codogno, P. (2000) Distinct classes of phosphatidylinositol 3′-kinases are involved in signaling pathways that control macroautophagy in HT-29 cells. J. Biol. Chem. 275,992–998.
Article CAS PubMed Google Scholar
Zeng, X., Overmeyer, J. H., and Maltese, W. A. (2006) Functional specificity of the mammalian Beclin-Vps34 PI 3-kinase complex in macroautophagy versus endocytosis and lysosomal enzyme trafficking. J. Cell. Sci. 119, 259–270.
Article CAS PubMed Google Scholar
Scarlatti, F., Bauvy, C., Ventruti, A., et al. (2004) Ceramide-mediated macroautophagy involves inhibition of protein kinase B and upregulation of beclin 1. J. Biol. Chem. 279, 18384–18391.
Article CAS PubMed Google Scholar
Sarkar, S., Floto, R. A., Berger, Z., et al. (2005) Lithium induces autophagy by inhibiting inositol monophosphatase. J. Cell Biol. 170, 1101–1111.
Article CAS PubMed Google Scholar
Criollo, A., Maiuri, M. C., Tasdemir, E., et al. (2007) Regulation of autophagy by the inositol trisphosphate receptor. Cell Death Diff., in press.
Google Scholar
Ogata, M., Hino, S. I., Saito, A., et al. (2006) Autophagy is activated for cell survival after ER stress. Mol. Cell Biol., in press.
Google Scholar
Kiffin, R., Bandyopadhyay, U., and Cuervo, A. M. (2006) Oxidative stress and autophagy. Antioxid. Redox Signal. 8, 152–162.
Article CAS PubMed Google Scholar
Kroemer, G., and Jaattela, M. (2005) Lysosomes and autophagy in cell death control. Nat. Rev. Cancer 5, 886–897.
Article CAS PubMed Google Scholar
Yoshimori, T., Yamamoto, A., Moriyama, Y., Futai, M., and Tashiro, Y. (1991) Bafilomycin A1, a specific inhibitor of vacuolar-type H(+)-ATPase, inhibits acidification and protein degradation in lysosomes of cultured cells. J. Biol. Chem. 266, 17707–17712.
CAS PubMed Google Scholar
Blommaart, E. F., Krause, U., Schellens, J. P., Vreeling-Sindelarova, H., and Meijer, A. J. (1997) The phosphatidylinositol 3-kinase inhibitors wortmannin and LY294002 inhibit autophagy in isolated rat hepatocytes. Eur. J. Biochem. 243, 240–246.
Article CAS PubMed Google Scholar
Xue, L., Borutaite, V., and Tolkovsky, A. M. (2002) Inhibition of mitochondrial permeability transition and release of cytochrome c by anti-apoptotic nucleoside analogues. Biochem. Pharmacol. 64, 441–449.
Article CAS PubMed Google Scholar
Moriyama, Y., and Nelson, N. (1989) H+-translocating ATPase in Golgi apparatus. Characterization as vacuolar H+-ATPase and its subunit structures. J. Biol. Chem. 264, 18445–18450.
CAS PubMed Google Scholar
Bampton, E. T., Goemans, C. G., Niranjan, D., Mizushima, N., and Tolkovsky, A. M. (2005) The dynamics of autophagy visualized in live cells: from autophagosome formation to fusion with endo/lysosomes. Autophagy 1, 23–36.
Article CAS PubMed Google Scholar
Kroemer, G., Galluzzi, L., and Brenner, C. (2007) Mitochondrial membrane permeabilization in cell death. Physiol. Rev. 87, 99–163.
Article CAS PubMed Google Scholar
Garrido, C., Galluzzi, L., Brunet, M., Puig, P. E., Didelot, C., and Kroemer, G. (2006) Mechanisms of cytochrome c release from mitochondria. Cell Death Differ. 13, 1423–1433.
Article CAS PubMed Google Scholar
Chernyak, B. V., Izyumov, D. S., Lyamzaev, K. G., et al. (2006) Production of reactive oxygen species in mitochondria of HeLa cells under oxidative stress. Biochim. Biophys. Acta 1757, 525–534.
Article CAS PubMed Google Scholar
Zamzami, N., Marchetti, P., Castedo, M., et al. (1995) Sequential reduction of mitochondrial transmembrane potential and generation of reactive oxygen species in early programmed cell death. J. Exp. Med. 182, 367–377.
Article CAS PubMed Google Scholar
Priault, M., Salin, B., Schaeffer, J., Vallette, F. M., di Rago, J. P., and Martinou, J. C. (2005) Impairing the bioenergetic status and the biogenesis of mitochondria triggers mitophagy in yeast. Cell Death Differ. 12, 1613–1621.
Article CAS PubMed Google Scholar
Matz, M. V., Fradkov, A. F., Labas, Y. A., et al. (1999) Fluorescent proteins from nonbioluminescent Anthozoa species. Nat. Biotechnol. 17, 969–973.
Article CAS PubMed Google Scholar
Rizzuto, R., Brini, M., Pizzo, P., Murgia, M., and Pozzan, T. (1995) Chimeric green fluorescent protein as a tool for visualizing subcellular organelles in living cells. Curr. Biol. 5, 635–642.
Article CAS PubMed Google Scholar
Rizzuto, R., Nakase, H., Darras, B., et al. (1989) A gene specifying subunit VIII of human cytochrome c oxidase is localized to chromosome 11 and is expressed in both muscle and non-muscle tissues. J. Biol. Chem. 264, 10595–10600.
CAS PubMed Google Scholar
Poot, M., Zhang, Y. Z., Kramer, J. A., et al. (1996) Analysis of mitochondrial morphology and function with novel fixable fluorescent stains. J. Histochem. Cytochem. 44, 1363–1372.
CAS PubMed Google Scholar
Daugas, E., Susin, S. A., Zamzami, N., et al. (2000) Mitochondrio-nuclear translocation of AIF in apoptosis and necrosis. FASEB J. 14, 729–739.
CAS PubMed Google Scholar
Galluzzi, L., Zamzami, N., De La Motte Rouge, T., Lemaire, C., Brenner, C., and Kroemer, G. (2007) Methods for the assessment of mitochondrial membrane permeabilization in apoptosis. Apoptosis, In press.
Google Scholar
Obeid, M., Tesniere, A., Ghiringhelli, F., et al. (2007) Calreticulin exposure dictates the immunogenicity of cancer cell death. Nat. Med., in press.
Google Scholar
Kroemer, G., and Reed, J. C. (2000) Mitochondrial control of cell death. Nat. Med. 6, 513–519.
Article CAS PubMed Google Scholar
Wei, M. C., Zong, W. X., Cheng, E. H., et al. (2001) Proapoptotic BAX and BAK: a requisite gateway to mitochondrial dysfunction and death. Science 292, 727–730.
Article CAS PubMed Google Scholar
Willis, S. N., and Adams, J. M. (2005) Life in the balance: how BH3-only proteins induce apoptosis. Curr. Opin. Cell Biol. 17, 617–625.
Article CAS PubMed Google Scholar
Zamzami, N., Larochette, N., and Kroemer, G. (2005) Mitochondrial permeability transition in apoptosis and necrosis. Cell Death Differ. 12 (Suppl. 2), 1478–1480.
Article CAS PubMed Google Scholar
Zoratti, M., and Szabo, I. (1995) The mitochondrial permeability transition. Biochim. Biophys. Acta 1241, 139–176.
PubMed Google Scholar
Cain, K., Bratton, S. B., and Cohen, G. M. (2002) The Apaf-1 apoptosome: a large caspase-activating complex. Biochimie 84, 203–214.
Article CAS PubMed Google Scholar
Martins, L. M., Iaccarino, I., Tenev, T., et al. (2002) The serine protease Omi/HtrA2 regulates apoptosis by binding XIAP through a reaper-like motif. J. Biol. Chem. 277, 439–444.
Article CAS PubMed Google Scholar
Du, C., Fang, M., Li, Y., Li, L., and Wang, X. (2000) Smac, a mitochondrial protein that promotes cytochrome c-dependent caspase activation by eliminating IAP inhibition. Cell 102, 33–42.
Article CAS PubMed Google Scholar
Verhagen, A. M., Ekert, P. G., Pakusch, M., et al. (2000) Identification of DIABLO, a mammalian protein that promotes apoptosis by binding to and antagonizing IAP proteins. Cell 102, 43–53.
Article CAS PubMed Google Scholar
Modjtahedi, N., Giordanetto, F., Madeo, F., and Kroemer, G. (2006) Apoptosis-inducing factor: vital and lethal. Trends Cell Biol. 16, 264–272.
Article CAS PubMed Google Scholar
Susin, S. A., Lorenzo, H. K., Zamzami, N., et al. (1999) Molecular characterization of mitochondrial apoptosis-inducing factor. Nature 397, 441–446.
Article CAS PubMed Google Scholar
Li, L. Y., Luo, X., and Wang, X. (2001) Endonuclease G is an apoptotic DNase when released from mitochondria. Nature 412, 95–99.
Article CAS PubMed Google Scholar
Metivier, D., Dallaporta, B., Zamzami, N., et al. (1998) Cytofluorometric detection of mitochondrial alterations in early CD95/Fas/APO-1-triggered apoptosis of Jurkat T lymphoma cells. Comparison of seven mitochondrion-specific fluorochromes. Immunol. Lett. 61, 157–163.
Article CAS PubMed Google Scholar
Ferri, K. F., Jacotot, E., Blanco, J., Este, J. A., and Kroemer, G. (2000) Mitochondrial control of cell death induced by HIV-1-encoded proteins. Ann. NY Acad. Sci. 926, 149–164.
Article CAS PubMed Google Scholar
Susin, S. A., Larochette, N., Geuskens, M., and Kroemer, G. (2000) Purification of mitochondria for apoptosis assays. Methods. Enzymol. 322, 205–208.
Article CAS PubMed Google Scholar
Goldstein, J., Waterhouse, N., Juin, P., Evan, G., and Green, D. (2000) The coordinate release of cytochrome c during apoptosis is rapid, complete and kinetically invariant. Nat. Cell Biol. 2, 156–162.
Article CAS PubMed Google Scholar
Loeffler, M., Daugas, E., Susin, S. A., et al. (2001) Dominant cell death induction by extramitochondrially targeted apoptosis-inducing factor. FASEB J. 15,758–767.
Google Scholar
Emoto, K., Toyama-Sorimachi, N., Karasuyama, H., Inoue, K., and Umeda, M. (1997) Exposure of phosphatidylethanolamine on the surface of apoptotic cells. Exp. Cell Res. 232, 430–434.
Article CAS PubMed Google Scholar
Martin, S. J., Reutelingsperger, C. P., McGahon, A. J., et al. (1995) Early redistribution of plasma membrane phosphatidylserine is a general feature of apoptosis regardless of the initiating stimulus: inhibition by overexpression of Bcl-2 and Abl. J. Exp. Med. 182, 1545–1556.
Article CAS PubMed Google Scholar
Rothe, G., and Valet, G. (1990) Flow cytometric analysis of respiratory burst activity in phagocytes with hydroethidine and 2’,7’-dichlorofluorescin. J. Leukoc. Biol. 47, 440–448.
CAS PubMed Google Scholar
Carter, W. O., Narayanan, P. K., and Robinson, J. P. (1994) Intracellular hydrogen peroxide and superoxide anion detection in endothelial cells. J. Leukoc. Biol. 55, 253–258.
CAS PubMed Google Scholar
Petit, P. X., Lecoeur, H., Zorn, E., Dauguet, C., Mignotte, B., and Gougeon, M. L. (1995) Alterations in mitochondrial structure and function are early events of dexamethasone-induced thymocyte apoptosis. J. Cell Biol. 130, 157–167.
Article CAS PubMed Google Scholar
Kroemer, G., and Martin, S. J. (2005) Caspase-independent cell death. Nat. Med. 11, 725–730.
Article PubMed Google Scholar
Garrido, C., and Kroemer, G. (2004) Life’s smile, death’s grin: vital functions of apoptosis-executing proteins. Curr. Opin. Cell Biol. 16, 639–646.
Article CAS PubMed Google Scholar
Fuentes-Prior, P., and Salvesen, G. S. (2004) The protein structures that shape caspase activity, specificity, activation and inhibition. Biochem. J. 384, 201–232.
Article CAS PubMed Google Scholar
Vahsen, N., Cande, C., Dupaigne, P., et al. (2006) Physical interaction of apoptosis-inducing factor with DNA and RNA. Oncogene 25, 1763–1774.
Article CAS PubMed Google Scholar
Niemann, A., Takatsuki, A., and Elsasser, H. P. (2000) The lysosomotropic agent monodansylcadaverine also acts as a solvent polarity probe. J. Histochem. Cytochem. 48, 251–258.
CAS PubMed Google Scholar
Harborth, J., Elbashir, S. M., Bechert, K., Tuschl, T., and Weber, K. (2001) Identification of essential genes in cultured mammalian cells using small interfering RNAs. J. Cell. Sci. 114, 4557–4565.
CAS PubMed Google Scholar
Nobukuni, T., Joaquin, M., Roccio, M., et al. (2005) Amino acids mediate mTOR/raptor signaling through activation of class 3 phosphatidylinositol 3OH-kinase. Proc. Natl. Acad. Sci. USA 102, 14238–14243.
Article CAS PubMed Google Scholar

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Authors and Affiliations

Unit “Apoptosis, Cancer and Immunity,”, INSERM, Villejuif, France
Ezgi Tasdemir, Lorenzo Galluzzi, M. Chiara Maiuri, Alfredo Criollo, Ilio Vitale, Emilie Hangen, Nazanine Modjtahedi & Guido Kroemer
Institut Gustave Roussy, Villejuif, France
Ezgi Tasdemir, Lorenzo Galluzzi, M. Chiara Maiuri, Alfredo Criollo, Ilio Vitale, Emilie Hangen, Nazanine Modjtahedi & Guido Kroemer
Faculté de Médecine–Université Paris-Sud, Villejuif, France
Ezgi Tasdemir, Lorenzo Galluzzi, Alfredo Criollo, Ilio Vitale, Emilie Hangen, Nazanine Modjtahedi & Guido Kroemer
Dipartimento di Farmacologia Sperimentale, Facoltà di Scienze Biotecnologiche–Università degli Studi di Napoli “Federico II”, Napoli, Italy
M. Chiara Maiuri

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Ezgi Tasdemir
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Lorenzo Galluzzi
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M. Chiara Maiuri
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Alfredo Criollo
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Ilio Vitale
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Emilie Hangen
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Nazanine Modjtahedi
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Guido Kroemer
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Department of Molecular Genetics and Microbiology, University of New Mexico Health Sciences Center, Albuquerque, NM, USA
Vojo Deretic

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Tasdemir, E. et al. (2008). Methods for Assessing Autophagy and Autophagic Cell Death. In: Deretic, V. (eds) Autophagosome and Phagosome. Methods in Molecular Biology™, vol 445. Humana Press. https://doi.org/10.1007/978-1-59745-157-4_3

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DOI: https://doi.org/10.1007/978-1-59745-157-4_3
Publisher Name: Humana Press
Print ISBN: 978-1-58829-853-9
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