In Vitro Assessment of Antioxidant Capacity, Potential to Induce Autophagy/Mitophagy, and Mitochondrial Turnover of Natural Compounds for Cytotoxicity Studies

Abstract

The dynamic process of intracellular reduction-oxidation status (redox homeostasis) is influenced by various factors, with mitochondria being one of the most significant contributors. Mitochondria play a crucial role in the bioenergetic pathway, fulfilling the metabolic energy demands of cells. To maintain increased energy requirements, mitochondrial biogenesis and fusion are employed, while decreased energy demands or damaged mitochondria are addressed through fission and autophagic removal, known as mitophagy. Any disruption in these adaptive responses can compromise redox homeostasis and cellular function, and make cells more vulnerable to oxidative stress, resulting in oxidative DNA damage, inflammatory responses, and apoptotic/anti-apoptotic reactions. Such dysregulation contributes to the development of “free radical diseases” like metabolic disorders and cancer. Traditional medicines and herbs (possessing antioxidant and autophagic properties) have been utilized for centuries in the treatment of various diseases; however, it is only recently that researchers have begun to investigate their molecular, cellular, and tissue-level modes of action. Nevertheless, concerns about their cytotoxicity have also arisen. This manuscript focuses on the current technological advancements in assessing the properties of plant-based natural compounds. Both cell-free and cell-based methods are employed to evaluate the therapeutic potential of these compounds, allowing for their scientific evaluation and validation.

1. Introduction

Past decades have perceived growing interest of studies investigating the role of Reactive Oxygen Species (ROS) and its amelioration with natural products [1, 2, 3, 4]. There are increasing evidences that suggest increased oxidative stress from ROS plays critical role in several pathophysiologies and causes Endoplasmic Reticulum stress and mitochondrial dysfunction. The intracellular redox balance (reduction–oxidation status) is a dynamic system which maintains the cellular homeostasis and may change via many factors [5, 6, 7]. Mitochondria, being one of the most important organelles, play a crucial role in maintaining intracellular source of energy. Any alteration in mitochondrial dynamics can cause an overproduction of ROS, which urges the induction of oxidative DNA damage and up- or down-regulation of phosphatases, proliferative/anti-proliferative factors, apoptotic/anti-apoptotic factors, etc. [5]. However, physiological levels of ROS are crucial for a proper cell function (i.e. intracellular signaling, inflammation, and immune function) [8]. Moreover, mitochondrial dysfunction and redox imbalance can contribute to a wide range of pathologies under a wide group termed as “free radical diseases” (e.g. cancer, neurodegeneration, atherosclerosis, inflammation, etc.) [5]. Altered cellular homeostasis by oncogenes, mitochondrial dysfunction, and/or oxidative stress leads to an increased genomic instability and the target cells can adapt to the changing environment [9].

Oxygen is vital for most unicellular and multicellular organisms, paradoxically it is also a key in generating Reactive Oxygen and Nitrogen Species (ROS/RNS). Endogenously, ROS are normal by-products of mitochondrial metabolism and energy production [5]. Exogenous sources include smoking, pollution, physical exercise etc. In metabolic disorders or cancer, ROS/RNS homeostasis compromises leading to oxidative stress [10, 11]. To maintain the fine balance and to counteract the harmful effects taking place in the cell, biological system has evolved itself with some strategies like prevention of oxidative damage, repair mechanism, finally the most important is the antioxidant defense mechanisms. Cells are protected from oxidative stress by several antioxidant mechanisms both inside and outside the mitochondria. These include manganese superoxide dismutase (Mn-SOD) in the mitochondria, copper/zinc-SOD in the cytosol, and extracellular-SOD in the interstitium [12]. Classically antioxidants were defined as molecules that inhibit or prevent oxidation of a substrate and are required to control any excess in these ROS/RNS and to maintain Redox homeostasis.

Plant-based natural products have been basis of treatment of many human diseases. The extracts of several plants have been used as therapeutic agents. Spices and herbs that are sources of natural antioxidants, can protect against oxidative stress and thus play preventive role in many diseases [13]. The medicinal properties of folk plants are mainly attributed to the presence of flavonoids, other organic, and inorganic compounds such as coumarins, phenolic acids, and antioxidant micronutrients and are able to counter various metabolic disorders, cancers, and COVID-19 [14, 15, 16, 17].

1.1 Role of mitochondrial oxidative stress in metabolic disorders and cancer

Metabolism refers to breaking down of food, in a highly-regulated manner, into simpler units of proteins, carbohydrates, and fats in order to generate energy for the organism. Metabolic syndrome is considered to be a progressive pathophysiological state which is clinically manifested by a cluster of interrelated risk factors such as abdominal obesity, atherogenic dyslipidemia, increased blood pressure, insulin resistance, and pro-inflammatory response [18, 19]. “Cancer metabolism” refers to the alterations in cellular metabolic pathways that are evident in cancer cells compared with most normal tissue cells. Major metabolic changes in cancer cells are increased glycolysis and reduced oxidative phosphorylation coupled with increased biosynthetic intermediates needed for cell growth and proliferation [20]. Likewise, the most common cellular patho-physiologies of metabolic disorders are increased oxidative stress, impaired mitochondrial functions, and cellular repair mechanism (autophagy and apoptosis), etc. [21, 22, 23].

Mitochondria are importantly placed in the history of cancer and have been attributed to play a crucial role in metabolic disorders [24, 25]. Although, mitochondria are the center for bioenergetics and biosynthesis pathways, they have also become a critical source of oxidative stress by generating ampule amount of superoxide in ETC [5]. Hence, mitochondrial redox control is utmost important for oxidative phosphorylation, ATP synthesis, and ROS production. The production of ROS in mitochondria is tightly regulated by the mitochondrial superoxide dismutase (SOD2) and glutathione-peroxidase (GPx), as well as by catalase and non-enzymatic antioxidants [5]. Excessive mitochondrial ROS and mitochondrial dysfunctions can promote metabolic−/age-related disorders (NAFLD, diabetes, atherosclerosis, neurodegeneration, etc.) and cancer through suppression of complex I, induction of oxidative mitochondrial DNA damage, increased excessive calcium ion influx, alterations in autophagy [5, 26, 27, 28]. Therefore, Phyto molecules/plant-based compounds having antioxidant potential and/or autophagic induction may provide beneficial effects in above-mentioned diseases that are associated with ROS and oxidative stress.

1.2 Therapeutic potential of natural compounds in metabolic disorders and cancer

Natural products have always been of immense importance in health and disease treatment throughout the human evolution [1, 2, 3, 4]. Traditionally several natural products have been used to treat different types of human illnesses including general injuries, wound healing, and pain [29, 30]. Natural products have been shown to have pharmacological properties to regulate cell signaling pathways that cause mitogenic and cytotoxic reactions leading to disease pathologies. Usage of modern technological tools such as genomics, proteomics, and metabolomics paved wider use of these natural products. Nowadays, mostly natural products are processed and have been developed as potential pharmacological agents with effective anti-oxidative, anti-mitotic, anti-microbial, anti-inflammatory, anti-angiogenic, and anti-carcinogenic properties [13, 31]. Different civilizations have been relying on plants or plant-based products to improve health. This also holds remarkable therapeutic alternatives to synthetic drugs. Millions of Europeans, Africans, Asians, Australians, and North Americans use plant-based medicine system as health supplements for their general health wellbeing. However, in developing countries, ayurveda and TCM, the traditional Indian and Chinese medicine is often the primary source of health care [32].

Many medicinal plants and natural products are considered by the public as a safe, natural, and cost-effective alternative to synthetic drugs without explicit proof by randomized controlled clinical trials [32]. Recently, there is an added attention in the validation of the efficacy and safety of natural products [32].

Several preclinical studies and non-scientific data accumulated over decades of research confirm the use of medicinal plants and plant-based products in the treatment or prevention of metabolic disorders. These products have gained popularity in western societies; however, their expected health benefits are often not backed by rigorous clinical trials. Therefore, it is essential to provide robust scientific evidence on their clinical efficacy and safety.

2. In vitro tools and techniques for the assessments of natural compounds

2.1 Assessment of antioxidant potential of plant extract/natural compound in a cell free system

2.1.1 Total antioxidant capacity by ABTS decolorization assay

Principle: The assay is based on the ability of antioxidant molecules to quench the stable ABTS (2,2´-azino-bis (3-ethylbenzthiazoline-6-sulfonic acid)) radical [33]. A blue-green solution of ABTS cation radical (ABTS^•+) is formed by the loss of an electron from stable ABTS radical and absorbs at 743 nm. In the assay, addition of Trolox decolorizes the solution resulted by the quenching the hydrogen atom by nitrogen atom.

Methodology (for 96-well plate format):

1. ABTS^•+ radical cation is produced by reacting ABTS^•+ stock solution (7 mM) with 2.45 mM potassium persulfate in the ratio of 1:0.5.

2. Allow the above-prepared mixture to stand in the dark at room temperature for 12–16 h.

3. Take 80 μl of ABTS solution prepared in step I and add 2.91 ml of ethanol to it.

4. Add 10 μl of plant extract (usually 1 mg/ml) or pure plant-based chemical and read it on a spectrophotometer immediately at 734 nm for 4–5 min.

5. 10 μl trolox was used as a standard. The decolorization of chromophore was measured on spectrophotometer.

6. Data should be presented as trolox equivalent antioxidant capacity (TEAC)/mg extract.

2.1.2 Nitric oxide (NO) quenching capacity

Principle: Nitric oxide (NO) interacted with oxygen to produce stable products, nitrite and nitrate that are key to generate RNS. Scavenger of NO competed with oxygen, leading to a reduced production of nitrite. The concentration of nitrite assays by the method of Feelisch and Noack [34] by reacting nitrite with griess reagent with which it forms a stable product which is measured at 542 nm on a spectrophotometer.

Methodology (for 96-well plate format):

1. Add 40 μl of phosphate buffer (20 mM, pH 7.4) to 10 μl of extract (1 mg/ml).

2. To the solution prepared in step I, add 50 μl of sodium nitroprusside (10 mM).

3. Incubate in dark at 37°C for 2.5 h.

4. Subsequently add 100 μl of griess reagent to it and after 30 min of incubation.

5. Read the product microplate reader at 542 nm.

6. Capacity of extract to inhibit this stable product is taken as NO quenching capacity/mg extract.

Application: Oxidative stress plays a key role on pathophysiology and aging [35, 36, 37]. Many phytochemicals/active compounds (polyphenols, flavonoids, terpenes, etc.) have antioxidant properties that are responsible for therapeutic effects of plants/herbs [14, 32]. Evaluating antioxidant potential of herbal/plant extract is the easy and very useful way to validate extract’s therapeutic efficacy toward oxidative stress.

2.2 Assessment of autophagy, lipophagy, and mitophagy using in vitro cell system

2.2.1 Analysis of autophagy by Western blotting

Introduction and principle: Autophagy is a catabolic cellular multistep process [38]. In autophagy, most commonly known as macroautophagy, targeted cytoplasmic constituents (such as damaged macromolecules and organelles) are isolated from the rest of the cell within a double-membraned vesicle called autophagosome. These autophagosome eventually fuses with lysosomes forming autolysosome and the contents inside are degraded under acidic environment and get recycled. Non-selective autophagy or macroautophagy is dissociated with other forms of specific autophagy processes such mitophagy (autophagic degradation of mitochondria), lipophagy (autophagic degradation of lipids), ERphagy (autophagic degradation of ER), etc.

Autophagy is executed by several autophagy-related (Atg) genes initially identified in Saccharomyces cerevisiae and later validated in mammalian systems [39]. During initiation of autophagy, two ubiquitin-like proteins Atg12 to Atg5 covalently conjugate and then bind to Atg16L1 to form an E3-like complex which functions as part of the second ubiquitin-like conjugation system. This complex binds and activates Atg3, which covalently attaches mammalian homologs of the ubiquitin-like protein Atg8/LC3B to the lipid phosphatidylethanolamine (PE) on the surface of autophagosomes after conversion from LC3B-I to LC3B-II. This lipidated LC3B (LC3B-II) closes the autophagosomes and enables docking of cargo-specific adaptor proteins such as Sequestosome-1/p62. The completed autophagosome then fuses with a lysosome through the actions of multiple proteins. After autophagosome-lysosome fusion, LC3B that is retained on the outer side is cleaved off and recycled whereas inside the vesicle along with Sequestosome-1/p62 gets degraded. Hence, Western blot analysis of the rate of accumulation of LC3B-II (autophagic flux) under lysosomal inhibition (using inhibitors such as chloroquine, Bafilomycin A1 etc.) is the most appropriate way to study autophagy and take as autophagic marker [40].

Methodology:

1. Seed the cells in appropriate density in multi-well plate and incubate at 370C and 5% CO2: 95% Air environment inside an incubator.

2. At 70–80% confluency, make four groups as i. Untreated; ii. Lysosomal inhibitor; iii. Treatment X (extract or natural compound); and iv. Treatment X + Lysosomal inhibitor.

3. Treat groups iii and iv with extract or natural compound for 24–72 h.

4. After the treatment is over, treat the groups ii and iv with Lysosomal inhibitor for 4–6 h for acute accumulation of autophagosomes.

5. Harvest cells for protein extraction using standard RIPA buffer and run western blot to analyze LC3B-II (14 kDa) and internal control (GADH).

Analysis of LC3B-II western blot: As shown in the (Figure 1A), there will be 3 cases of LC3B-II western blot image.

Figure 1.

Analysis of autophagic flux by Western blotting.

Step-I: Take normalized densitometric ratios for basal autophagy (yellow box): i.e. lanes 2:1.

Step-II: Take normalized densitometric ratios for autophagy under treatment (green box): i.e. lanes 4:3.

Step-III: Compare the ratios from basal autophagy (step-I) vs. autophagy under treatment (step-II) for significance.

Interpretation of LC3B-II western blot:

As mentioned earlier that autophagy is a kinetic process. However, during the analysis, the expression of cellular LC3B-II at a given time point is a static and unable to represent the rate of autophagosome synthesis and degradation after lysosomal fusion. i.e. autophagy flux. Thus, to better analyze this, use of lysosomal inhibitors is important. This enables us to analyze any increase in the LC3B-II in the presence of a lysosomal inhibitor representing autophagy induction or autophagosome biogenesis before lysosomal activity was blocked. Hence, as shown in the Figure 1A, there will be 3 cases (Figure 1B–D) of autophagy flux according to LC3B-II expression before and after the treatment of lysosomal inhibitors.

• Case-1: Activation of autophagic flux by treatment X.

If relative LC3B-II protein expression is higher under treatment when compared to untreated group (as shown in Figure 1B by red dotted line), and it is further increased after lysosomal inhibition (as shown in Figure B by double-head arrows) in treated group compared to untreated group.

• Case-2: Inhibition of autophagic flux by treatment X at autophagosome formation step (i.e. early block).

If relative LC3B-II protein expression is lower under treatment when compared to untreated group (as shown in Figure 1B by red dotted line), and it is further decreased or unchanged after lysosomal inhibition (as shown in Figure 2B by double-head arrows) in treated group compared to untreated group.

• Case-3: Inhibition of autophagic flux by treatment X at autophagosome maturation/autolysosome step (i.e. late block).

• If relative LC3B-II protein expression is higher under treatment when compared to untreated group (as shown in Figure 1D by red dotted line; and this can be easily miss-interpreted to autophagy activation without flux analysis), and it is decreased or unchanged after lysosomal inhibition (as shown in Figure 1D by double-head arrows) in treated group compared to untreated group.

2.2.2 Analysis of autophagy and mitophagy by fluorescence microscopy

Introduction and principle: To analyze the progression of autophagy/mitophagy from autophagome/mitophagosomes to lysosomes, recently developed reporter plasmids (Figure 2), that are tandem-tagged RFP-EGFP protein fused and cloned with LC3B (to analyze autophagy) or mitochondrial localization signal (MLS) typically COX-IV (also called mitoRFP-GFP plasmid; to analyze mitophagy), have been used, which exploit the differential stabilities of RFP and GFP in an acidic environment [40]. Green GFP signals are quenched in acidic pH, whereas RFP signals can be visualized in both autophagosome and acidic autophagolysosome, thus an increase in RFP fluorescence in the lysosomes indicates completion of the mitophagy process. However, at basal, yellow signals indicate merged images of GFP and RFP signals for only autophagosome containing mitochondria. Therefore, the formation of red puncta in treatment groups will indicate increased autophagic/mitophagic flux over untreated cells which will show yellow puncta.

Figure 2.

RFP-GFP fusion plasmids to study autophagic/mitophagic flux under fluorescent microscope.

Methodology:

1. Seed the cells in appropriate density in chambered slides and incubate at 370C and 5% CO2: 95% Air environment inside an incubator.

2. Plasmid transfection: At 70–80% confluency, make two groups as i. untreated and ii. Treatment X (extract or natural compound), then use 0.5 μg plasmid and transfection reagent (usually Lipofectamin P3000 from Invitrogen, USA) to transfect DNA into the cells from both the groups, and incubate for 24 h at 370C and 5% CO2: 95% Air environment inside an incubator.

3. Treat group ii with extract or natural compound for further 24–48 h.

4. After the treatment is over, fix the cells in 4% paraformaldehyde for 15 min.

5. Stain cells with Hoechst 33342 or DAPI (Sigma) to stain nucleus.

6. Wash the cells with 1x phosphate buffer saline (PBS) two times for 5 min each.

7. Mount the slide using anti-fade reagent and visualize under the fluorescent microscope or confocal microscope at higher magnifications (40–60x is recommended).

Analysis of fluorescent images: Both RFP and GFP fluoresces on autophagosomes as LC3B and represents as yellow puncta in overlay under basal condition. However, in autolysosome, under acidic/low pH GFP (green) fluorescence get quenched and only RFP fluorescence can be visualized as red puncta (in overlay) of autolysosome suggesting increase in autophagy flux. Similarly, both RFP and GFP fluoresces on mitochondria and whole mitochondrial network can be visualized under basal condition. However, in mitophagolysosome, under acidic/low pH GFP (green) fluorescence get quenched and only RFP fluorescence will be visualized as red puncta (in overlay) of mitophagolysosome suggesting increase in mitophagy flux.

2.2.3 Analysis of lipophagy by fluorescence microscopy

Introduction and principle: To analyze lipophagy (autophagic degradation of lipids), cellular lipid droplets/triglycerides stain using BODIPY™ 493/503 dye (Invitrogen) and endogenous LC3B proteins stain using anti-LC3B antibody. Fluorescence microscopy analysis is performed to analyze co-localization of lipids/BODIPY with LC3B.

Methodology:

1. Seed the cells in appropriate density in chambered slides and incubate at 370C and 5% CO2: 95% Air environment inside an incubator.

2. At 70–80% confluency, make two groups as i. untreated and ii. Treatment X (extract or natural compound), treat group ii with extract or natural compound for further 24–48 h, and incubate at 370C and 5% CO2: 95% Air environment inside an incubator.

3. After the treatment is over, fix the cells in 4% paraformaldehyde for 15 min.

4. Wash cells with 1x phosphate buffer saline (PBS) three times 5 min each.

5. Incubate cells with 1% BSA in PBST (PBS + 0.1% Tween 20) + 10% serum from the species the secondary antibody was raised in, for 1 h at room temperature to block unspecific binding of the primary antibody.

6. Remove blocking reagent carefully and incubate cells in the diluted primary antibody (anti-LC3B) in 1% BSA in PBST overnight at 4°C.

7. Wash cells with 1x phosphate buffer saline (PBS) three times for 5 min each.

8. Incubate cells with the secondary antibody (Alexa Fluor® 594; Invitrogen) in 1% BSA in PBST for 1 h at room temperature in the dark.

9. Remove reagent, wash one time in 1x PBS and incubate cells in BODIPY™ 493/503 stain (1:5000 dilution) for 15 min at room temperature.

10. Stain cells with Hoechst 33342 or DAPI (Sigma) to stain nucleus.

11. Wash cells with 1x phosphate buffer saline (PBS) twice for 5 min each.

12. Mount the slide using anti-fade reagent and visualize under the fluorescent microscope or confocal microscope at higher magnifications (40–60x is recommended).

Analysis of fluorescent images: Both BODIPY™ fluorescence is visualized using GFP/Annexin filter to observe green fluorescence of lipid droplets, however, RFP filter can be used to visualize LC3B red fluorescence. Lipophagy will be seen in overlay as yellow puncta formed by BODIPY-LC3B co-localization.

2.2.4 Analysis of autophagy, lipophagy, and mitophagy by electron microscopy

Introduction and principle: Electron microscopy is one of the best methods for the detection of autophagy and quantification of autophagic accumulation compared to fluorescence microscopy that has lower resolution than transmission electron microscopy [41]. During autophagy induction, the number of degradative compartments increases and the quantification of the number of autophagosome, autolysosomes, lysosomes, and amphisomes per cell section provides a simple morphological readout. But, it is very difficult to distinguish between autolysosomes, lysosomes, and amphisomes, hence, it is much simpler to classify all these organelles together, as degradative compartments or autophagic vesicles. However, autophagosome and autolysosome can be distinguished by observing their double membrane vs. single membrane walls respectively.

Methodology:

1. Seed the cells in appropriate density in chambered glass coverslips and incubate at 370C and 5% CO2: 95% Air environment inside an incubator.

2. At 70–80% confluency, make two groups as i. untreated and ii. Treatment X (extract or natural compound), treat group ii with extract or natural compound for further 24–48 h, and incubate at 370C and 5% CO2: 95% Air environment inside an incubator.

3. Fix the monolayers in 2% glutaraldehyde (EM grade) in 0.1 M phosphate buffer pH 7.4 for 1 h at room temperature.

4. Wash with phosphate buffer twice for 3 min each.

5. Mix osmium tetroxide and NaCac in water to give final concentrations of 1% and 0.1 M respectively. Next, add potassium ferrocyanide (K4Fe(CN)6) at 15 mg/ml, add on to the cells and incubate for 1 h at room temperature.

6. Wash with phosphate buffer twice for 3 min each, then with water once.

7. Incubate the monolayers in 1% uranyl acetate and 0.3 M sucrose in water for 1 h at 4°C.

8. Dehydrate cells with 70% ethanol for 1 min followed by 96% ethanol for 1 min, and 100% ethanol for 1 min three times.

9. Dip cover slips into acetone and immediately drop some epoxy resin to cover the cells, before the acetone has completely evaporated.

10. Fill beem capsules with resin and place them upside down on the cover slips. Incubate for 2 h at room temperature and let the resin polymerize at 60°C overnight.

11. Carefully remove the resin block from the cover slips and trim the block. Cut sections of 70- to 80-nm thickness.

12. Post-stain the sections with uranyl acetate and lead citrate for 3 min for grid preparation [42].

13. Grids can be visualized under electron microscope and images can be acquired at 10000x as primary magnification, and 40,000x to 100,000x magnification can be used to visualized autophagic vesicles containing cargo (lipid droplets, mitochondria, etc.).

Application: Autophagy plays a key role in the cellular maintenance and development of disease [38, 43]. Impairment in autophagic removal of damaged mitochondria (mitophagy) and fatty acids/lipids (lipophagy) has been shown to be important in various pathological states including neurodegenerative diseases and metabolic disorders [38, 43]. Recently, phytochemicals have been shown to be an autophagy inducers and are beneficial in various diseases and metabolic disorders [44, 45, 46, 47]. Hence, analyzing autophagy flux and their potential to remove damaged mitochondria and lipids can be an easy way to evaluate plant’s therapeutic efficacy.

2.3 Assessment of mitochondrial turnover (synthesis of new and removal of damaged mitochondria) using MitoTimer

Introduction and principle: Fluorescent Timer, or DsRed, is a mutant of the red fluorescent protein, dsRed, in which fluorescence shifts over time from green to red as the protein matures or undergoes oxidation during the course of aging. This molecular clock gives temporal and spatial information on protein turnover [48]. To monitor mitochondrial turnover, this Timer is engineered and tagged with mitochondrial-targeting sequence (called as “MitoTimer”) and cloned in a plasmid. When cells get MitoTimer, it localizes in the mitochondria and disappearance of red fluorescence (for removal of damaged mitochondria) and appearance of green mitochondrial (for synthesis of new mitochondria) i.e. turnover, can be visualized.

Methodology:

1. Seed the cells in appropriate density in chambered slides and incubate at 370C and 5% CO2: 95% Air environment inside an incubator.

2. Plasmid transfection: At 70–80% confluency, make two groups as i. untreated and ii. Treatment X (extract or natural compound), then use 0.5 μg plasmid and transfection reagent (usually Lipofectamin P3000 from Invitrogen, USA) to transfect DNA into the cells from both the groups, and incubate for 24 h at 370C and 5% CO2: 95% Air environment inside an incubator.

3. Treat group ii with extract or natural compound for further 24–48 h.

4. After the treatment is over, fix the cells in 4% paraformaldehyde for 15 min.

5. Stain cells with Hoechst 33342 or DAPI (Sigma) to stain nucleus.

6. Wash the cells with 1x phosphate buffer saline (PBS) two times for 5 min each.

7. Mount the slide using anti-fade reagent and visualize under the fluorescent microscope or confocal microscope at higher magnifications (40–60x is recommended).

Analysis of fluorescent images: As shown in the [49], both mitoRFP and mitoGFP fluoresces on mitochondria and represented as yellow mitochondrial network in overlay under basal condition, suggesting mitochondrial homeostasis. However, when there is an increase in mitochondrial turnover by mitophagy, mitoGFP (green) fluorescence increases and mitoRFP fluorescence decreases under (A) treatment X suggesting increase in new mitochondria via biogenesis and removal of old/damaged mitochondria by mitophagy. Similarly, when turnover is decreased, both mitoRFP increases and mitoGFP decreases suggesting decreased mitochondrial biogenesis and accumulation of damaged/aged mitochondria under (B) treatment X.

Application: Mitochondrial turnover is utmost important to maintain mitochondrial homeostasis that is required for cellular health [5, 50]. Dysregulated mitochondrial turnover leads to cause oxidative stress and cell death which is a major cause for metabolic disorders and cancer [24, 26, 51]. Study of mitochondrial turnover using MitoTIMER assay is a convenient method to evaluate phytochemicals efficacy.

2.4 Assessment of cell viability/cytotoxicity in vitro

2.4.1 MTT tetrazolium assay

Principle: Tetrazolium salts are widely used for detecting redox potential of cells for viability, cytotoxicity, and proliferation assays. This is a colorimetric assay that measures the reduction of yellow 3-(4,5-dimethythiazol2-yl)-2,5-diphenyl tetrazolium bromide (MTT) by mitochondrial succinate dehydrogenase, which demonstrates functional mitochondria [52]. The MTT is cell permeable and goes into the mitochondria and gets reduced to purple colored formazan crystals. These purple crystals are then solubilized using organic solvent. Lastly, this purple solution is measured spectrophotometrically. Notably, reduction of MTT only occurs in metabolically active cells, thus, the level of purple color is a measure of the viable cells.

Methodology:

1. Seed the cells in appropriate density in a 96-well plate and incubate at 370C and 5% CO2: 95% Air environment inside an incubator.

2. At 70–80% confluency, make two groups as i. untreated and ii. Treatment X (extract or natural compound), treat group ii with extract or natural compound for further 24–48 h, and incubate at 370C and 5% CO2: 95% Air environment inside an incubator.

3. After treatment, add 20 μl /well of MTT solution (Sigma) and incubate for 4–5 h in 37oC in dark.

4. Aspirate MTT containing media and add 200 μl Di methyl Sulphoxide (DMSO, Sigma), mix gently and incubate for 1 h in 37oC in dark.

5. Measure absorbance at 570 nm (and background wavelength is 630 nm) using 96-well plate spectrophotometer.

6. The linear relationship between absorbance and cell number should be taken into account and the data should be expressed as a percentage of viability in untreated group.

2.4.2 MTS assay

Principle: Colorimetric method for sensitive quantification of viable cells in proliferation and cytotoxicity assay. This assay is also based on the same principle as MTT assay, where we read colored formazan product by metabolically active cells is soluble in cell culture media. The MTS assay offers a number of advantages over the MTT assay. First, it eliminates the need for washing or solubilization steps, making it a more efficient and streamlined process. This high-throughput assay can be performed directly in the cell culture media, and can be conducted using a 96-well microtiter plate. The MTS assay is versatile and can be used to measure cell proliferation in response to a variety of stimuli, such as growth factors, cytokines, mitogens, and nutrients. Additionally, it is useful for analyzing the cytotoxic effects of compounds such as anticancer drugs, as well as other toxic agents, and pharmaceuticals.

Methodology:

1. Seed the cells in appropriate density in a 96-well plate and incubate at 370C and 5% CO2: 95% Air environment inside an incubator.

2. At 70–80% confluency, make two groups as i. untreated and ii. Treatment X (extract or natural compound), treat group ii with extract or natural compound for further 24–48 h, and incubate at 370C and 5% CO2: 95% Air environment inside an incubator.

3. Add 20 μl/well MTS Reagent into each well and incubate for 4 h at 37°C in standard culture conditions.

4. Shake the plate briefly on a shaker and measure absorbance at 490 nm using 96-well plate spectrophotometer.

5. The linear relationship between absorbance and cell number should be taken into account and the data should be expressed as a percentage of viability of untreated group.

2.4.3 Luciferase-based assay for real-time cytotoxicity analysis

Principle: The latest innovation in viable cell number measurement is a real-time approach using a pro-substrate and luciferase. This method involves adding the reagent directly to the culture medium, where viable cells with active metabolism reduce the pro-substrate into a Luc-substrate. The Luc-substrate then diffuses into the culture medium, and luciferase utilizes it to generate a luminescent signal.

The pro-substrate is non-reactive with luciferase and is well-tolerated by cells, remaining stable in complete cell culture medium at 37°C for at least 72 hours. This long-term stability allows for multiple measurements from the same sample over a period of days without the need for pro-substrate replenishment.

The assay can be performed in both continuous-read and endpoint formats. In continuous-read mode, the luminescent signal is recorded repeatedly from the same sample wells over an extended period to measure the number of viable cells in real time. This new approach provides a reliable and efficient way to monitor viable cell numbers and metabolic activity in a variety of research applications [53].

Methodology:

RealTime-GloTM MT Cell Viability Assay (Promega) for continuous read mode:

1. Equilibrate the MT Cell Viability Substrate and the NanoLuc® Enzyme to 37°C.

2. Seed the cells in appropriate density in a white colored 96-well plate optimize for luminescence.

3. Add MT Cell Viability Substrate and the NanoLuc® Enzyme to 1X final concentration to the cell suspension and incubate at 370C and 5% CO2: 95% Air environment inside an incubator.

4. After 24 h, make two groups as i. untreated and ii. Treatment X (extract or natural compound), treat group ii with extract or natural compound.

5. Measure luminescence using a 96-well plate luminometer, and return cells to the cell culture incubator between measurements to maintain a consistent temperature at each measurement. To determine the linear range of cell densities for the continuous-read format, continue measuring luminescence at various times over the desired time course (e.g. 12 h, 24 h, 72 h). The data should be expressed as a percentage of initial viability (at 0 h) of respective groups.

Application: Analysis of cell viability plays a diverse role in establishing the relationship between drug safety and its toxicity. High-throughput methods are very well embraced in the scientific community for drug’s safety evaluation tastings. Hence, above-mentioned cytotoxicity/cell viability assays are best fit to titrate the safe dosage of phytochemicals.

3. Discussion

One of the important and attractive uses of the in vitro methods is to evaluate therapeutic efficacy and safety of active compounds/phytochemicals and/or crude extracts in a high-throughput manner, and that also limits the use of animals which can be expensive, labor-intensive, and time taking. The most common assays (as mentioned above in Section 2.1) are to evaluate antioxidant potential of herbal extracts or active compounds in a cell-free system or by using animal cell culture system. Prof. Y. Osumi has received Noble prize for re-stating the role of autophagy in cell health and disease [54]. Technological advancements in scientific field further made it possible to evaluate autophagic flux and related processes, such as mitophagy and lipophagy, in a high-throughput way by using fusion proteins (as discussed above in Sections 2.2 and 2.4) in animal cell culture system. Mitochondrial dysfunction and de-regulation of mitochondrial turnover and homeostasis play key roles in may diseases including cancer. Analysis of OCR that is an indicator of mitochondrial respiration and mitochondrial activity is utmost important for safety evaluation plant-based therapeutic compounds. Recently developed engineered mitochondria-localized fluorescent proteins [48], that aged or oxidized in time-dependent manner and fluoresce differentially (as discussed above in Section 2.3) are boon for new age research, can be used to evaluate mitochondrial turnover in a high-throughput way. Cytotoxicity is desirable in the cancer-targeted drugs; however, it can be detrimental for normal cells, so analysis of cytotoxicity is important for successful use of plant-based chemo-drugs or to target metabolic diseases. Although, there are several high-throughput assays available for assessing the potential cell viability and cytotoxicity of plant extracts and phytochemicals, this chapter (in Section 2.5) provides a summary of both commonly used and recently developed cell-free and cell-based assays. These assays serve as important tools for evaluating the safety and therapeutic effectiveness of plant-based drugs and compounds.

Acknowledgments

The authors are thankful to the Ministry of Health (MOH), and National Medical Research Council (NMRC), Singapore, for financial assistance grant number NMRC/OFYIRG/0002/2016 and MOH-000319 (MOH-OFIRG19may-0002) to BKS; NMRC/OFYIRG/077/2018 to MT; and CSAI19may-0002 to PMY.

Conflict of interest

The authors declare no conflict of interest.

Notes/thanks/other declarations

None.