Roots of Asparagus racemosus Willd. were collected from authenticated sources and graded accordingly. Voucher specimens were deposited as Asparagus racemosus Willd. (AMAR-1) at the Botanical Survey of India, Koregaon Road, Pune, and authenticated with certificate number No.BSI/VWRC/lden.Cer./2024/0901240015667. The phytoextraction processes, with minor adjustments to established protocols, were utilized for the hydroalcoholic (HAAR) and aqueous (AQAR) extraction of Asparagus racemosus (Rodríguez-García et al., 2019; Huei et al., 2020; Somaida et al., 2020; Borse et al., 2021; El-Desouky, 2021; Jadaun et al., 2023b). Briefly, the roots of Asparagus racemosus were first minced using a mortar and pestle, followed by further grinding with a mixer to obtain a fine powdered consistency. For the hydroalcoholic extract of Asparagus racemosus roots, powdered raw materials were soaked in a 70:30 water-alcohol mixture at a 1:4 ratio (material) overnight. The extraction process was conducted for 3 h at a temperature of 60 ± 5°C. The resulting extract was filtered through a 400-micron strainer three times to ensure purity. The filtered extract was then pooled, concentrated, and subjected to spray drying to obtain a final dry powder. For the aqueous extract, crushed Asparagus racemosus root powder was macerated in 3 liters of UV-filtered water for 8 h overnight. After maceration, the mixture was gently boiled for up to 8 h to reduce the volume to 1/8th of its original amount. The resulting concentrated decoction was then filtered using a cotton cloth to remove any solid residues. Finally, the filtrate was lyophilized to obtain a dry powder extract. To prepare stock solutions for both AQAR and HAAR, the aqueous extract was dissolved in water, while the hydroalcoholic extract was dissolved in DMSO. For subsequent assays, the stock solutions were diluted in complete medium or PBS. Care was taken to ensure that the final concentrations of alcohol and DMSO were minimal and maintained within non-toxic levels for the cells. The presence of Shatavarin IV in the root extracts HAAR and AQAR was previously confirmed by our research group (Borse et al., 2021), and reaffirmed in the current study using a High-Resolution Mass Spectrometry (HR-MS). HR-MS (ESI) analysis revealed peaks at m/z calcd for C₄₅H₇₄O₁₇ [M + H] + 887.4990 for AQAR and m/z calcd for C₄₅H₇₄O₁₇ [M + H] + 887.5001 for HAAR, with corresponding found values of 887.4999 for both extracts. These results further confirmed the presence of Shatavarin IV in both AQAR and HAAR extracts (Supplementary Figure S1). The reference pure compound Shatavarin IV was procured commercially from Sigma-Aldrich Solution, Merck, Darmstadt, Germany (CAS Number: 84633-34-1) for the in vitro evaluation of anti-HIV-1 activity in this study.

TZM-bl cells (HeLa modified cell line; initially called JC53-bl; clone 13) were acquired from the National Institute of Health (NIH)–HIV Reagent Program and maintained in DMEM (Gibco, MA, United States) supplemented with 10% FBS (Moregate, Bulimba, QLD, Australia), HEPES (Gibco, Waltham, MA, United States), and antibiotics (Sigma-Aldrich, St. Louis, MO, United States) at 37°C in a 5% CO₂ humidified chamber. Cells with at least 80% confluency were utilized for subsequent experiments. Peripheral blood mononuclear cells (PBMCs) were isolated from healthy individuals’ blood, collected from the discarded blood bags received from the government blood bank, using density gradient centrifugation with Histopaque (Sigma-Aldrich, St. Louis, MO, United States) and activated with PHA-P (5 μg/mL) (Sigma-Aldrich, St. Louis, MO, United States) in RPMI 1640 medium (Gibco, Waltham, MA, United States) supplemented with 10% FBS and 5 U/mL Interleukin-2 (IL-2) (Sigma-Aldrich, St. Louis, MO, United States) for growth factor support. Activated PBMCs were employed for HIV-1 stock generation and confirmation of anti-HIV-1 activity.

The HIV-1_UG070 (X4, Subtype D), a primary isolate, was obtained from NIH ARRRP, while HIV-1_VB28 (R5, Subtype C), Indian primary isolate, was grown at the Division of Virology, ICMR-NITVAR. Virus stocks were prepared in PHA-P (5 μg/mL) activated PBMCs derived from healthy donors and quantified by HIV-1 p24 antigen detection assay (Abcam, Cambridge, United Kingdom). These virus stocks were titrated in TZM-bl cell lines, and the TCID₅₀ (i.e., 50% of the tissue culture infective dose) was determined using the Spearman-Karber method.

Cellular toxicity profiles of the AR extracts and Shatavarin IV were determined using the MTT assay with TZM-bl and activated PBMCs. TZM-bl cells were seeded in 96-well plates and allowed to adhere overnight, while PBMCs were seeded directly in suspension. Serial dilutions of the extracts (0.0078–1.00 mg/mL) and/or the active molecule (0.039–5.00 mg/mL) were added to the cells, and the MTT Assay was performed after 48 h of incubation. Cell viability was measured colorimetrically, and the percentage of cell viability was calculated. The assay indirectly measured cellular toxicity, and the results were expressed as CC₅₀, the concentration of a product with at least 50% viable cells. Additionally, the ATPlite assay (PerkinElmer, Netherlands) was employed for the assessment of cell cytotoxicity. This assay is a highly sensitive and widely used method for determining cell viability and cytotoxicity. It is based on the quantification of adenosine triphosphate (ATP), which is present in all metabolically active cells. In this assay, cells were seeded in 96-well plates and treated with varying concentrations of AR extracts or Shatavarin IV. Following incubation, the ATPlite reagent was added to each well, and luminescence was measured using a multi-mode microplate reader (Perkin Elmer, United States). The luminescence signal was directly proportional to the amount of ATP present, allowing for the determination of cell viability. The concentration at which 50% of cell viability was inhibited (CC₅₀) was calculated from dose–response curves constructed from the data obtained. Both MTT and ATPlite assays were performed in triplicates.

Subtoxic concentrations of AQAR and HAAR extracts and/or bioactive constituent Shatavarin IV were tested against HIV-1 primary isolates using the cell-associated anti-HIV-1 assay. TZM-bl cells (1 × 10⁴ cells/well) were seeded in microplates prior to the assay. Pre-titrated HIV-1_VB028 and HIV-1_UG070 virus stocks were used to infect TZM-bl cells. Two-fold serial dilutions of the drug candidates were overlaid onto the infected cells, and luciferase activity was measured after 48 h of incubation using the Britelite plus reagent (Perkin Elmer, Waltham, MA, United States). A known reverse transcriptase inhibitor, azidothymidine or AZT, was taken as a standard drug control. Dose-dependent percent inhibition was calculated using the formula below: Percentage Inhibition = 1 − A v g R L U Test − A v g R L U Mock A v g R L U V C − A v g R L U Mock × 100

The effective inhibitory concentration for 50% inhibition of HIV-1 replication (EC₅₀) and for 80% inhibition of HIV-1 replication (EC₈₀) values were calculated. The reproducibility of results was confirmed by performing three independent assays.

To confirm the safety and efficacy profiles of AQAR, HAAR, and their bioactive molecule Shatavarin IV, following the initial screening using TZM-bl cells, confirmatory experiments were conducted. These experiments employed peripheral blood mononuclear cells (PBMCs), the natural host cells of HIV. PBMCs were utilized post-activation with PHA-P (5 μg/mL) to ensure their susceptibility to HIV infection. The activated cells were cultured in RPMI supplemented with 10% FBS, 25 mM HEPES buffer, Penicillin & Streptomycin (50 U/mL & 50 mg/mL), and 10 U/mL of IL-2 as described earlier. After 48 h of activation, the cells were washed and resuspended in RPMI supplemented with 2% FBS and seeded in 96-well plates (0.2× 10⁴ cells/well) prior to infection with HIV-1_VB028 (40 TCID₅₀). The infected PBMCs were treated with non-toxic concentrations of AR extracts and/or Shatavarin IV, and the experimental controls, including mock-treated cells (Mock), virus-infected cells (VC), and cells treated with a standard drug AZT (SD), were maintained throughout the assay. The seeded microplates were incubated for 5 days, and culture supernatants were harvested by centrifugation. HIV-1 p24 antigen levels in the supernatants were quantified using an ELISA kit following the manufacturer’s instructions (Abcam, Cambridge, United Kingdom). The assay was performed in triplicates, and percent inhibition along with EC50 values were calculated. The results were compared to the reference drug AZT.

To determine the viral copy number, activated PBMCs were employed. These PBMCs were infected with pre-titrated HIV-1_VB028 (40 TCID₅₀) strains for 4 h under conditions similar to those mentioned above. Subsequently, post-infection, the cells underwent triple washing with RPMI medium (Gibco, MA, United States) containing 2% FBS (Moregate, Bulimba, QLD, Australia). Following this, the cells were seeded into six-well plates and suspended in RPMI containing 10% FBS media. HAAR and AQAR extracts, along with Shatavarin IV, were added. On the eleventh day post-infection, the wells were terminated, and the HIV-1 viral copy number was assessed using the Abbott Real-Time Platform (Abbott, Chicago, IL, United States). The viral load assay was performed in triplicates to ensure reliability and reproducibility, and the results were compared with the reference drug AZT.

The inhibitory effects of AQAR and HAAR on HIV-1 integrase (INT) activity were assessed using a commercially available HIV-1 Integrase Assay Kit (XpressBio, Frederick, MD, United States) following the manufacturer’s instructions. Briefly, a Streptavidin-coated 96-well plate was coated with a double-stranded HIV-1 LTR U5 donor substrate (DS) oligonucleotide labelled with biotin. Recombinant HIV-1 integrase protein was then loaded onto the DS DNA substrate. Following the addition of AR extracts, a new double-stranded target substrate (TS) DNA with a 3′-end alteration was introduced into the enzyme reaction. Positive controls included 1.0% Sodium Azide (provided with the kit PC) and the known HIV-1 integrase inhibitor RAL (0.48 μM – Standard Drug or SD), while the integrase enzyme provided with the kit served as the negative control (Enzyme Control or EC). The activity of HIV-1 integrase involves cleaving the last two bases from the exposed 3′ end of the HIV-1 LTR DS DNA, which then integrates into the TS DNA through a strand-transfer recombination process. The reaction products were detected calorimetrically using an HRP-labelled antibody directed against the TS 3′-end alteration, and the absorbance resulting from the HRP antibody-TMB peroxidase substrate reaction was measured at 450 nm. Three independent assays were performed to ensure accuracy and consistency of the results.

The AR extracts were assessed for potential HIV-1 Protease inhibitory activity using an HIV-1 PR inhibitor screening Fluorometric assay kit (Abcam, Cambridge, United Kingdom). Each sample was incubated with the HIV-1 PR enzyme for 15 min at room temperature. Subsequently, the fluorescent substrate was added to the wells, and absorbance was measured (excitation/emission = 330/450 nm) using a plate reader in kinetic mode for 120 min at 37°C. The kit-supplied Enzyme Control (EC) served as the negative control, while the Inhibitor Control (PC) containing Pepstatin (1 mM) and known Protease inhibitor RTV (10 μM – SD) acted as positive controls to assess the inhibition of HIV-1 Protease activity. DMSO (1%, v/v) was utilized as the vehicle to normalize background noise. Assays were replicated three times for the reliability. The percentage inhibition of HIV-1 Protease was calculated based on the Relative Fluorescence Unit (RFU) of each test sample.

The inhibitory effects of AQAR and HAAR on HIV-1 Reverse Transcriptase (RTase) were assessed using a commercially available kit (Roche, Penzberg, Germany). In this assay, the extracts at their respective EC₈₀ concentrations were incubated with HIV-1 RTase and a template nucleotide mixture for 1 h. The resulting mixture was then transferred to streptavidin-coated microwell plates, where it bound to biotin, forming a complex with the DIG-labelled template primer. Subsequently, HRP-conjugated enzyme was added to each well, followed by another incubation period of 1 h. Absorbance readings were taken at wavelengths of 405 nm and 490 nm using a BioRad reader PR4100 after the addition of the substrate. Three independent assay replicates were taken into consideration for the data interpretation. AZT, a known inhibitor of HIV-1 RTase, served as a positive control (SD) in the assay.

The crystal structures of HIV-1 proteins, including HIV-1 Integrase (PDB: 1QS4, resolution 2.01 Å), Protease (PDB: 5KR0, resolution 1.8 Å), and Reverse Transcriptase (PDB: 3QIP, resolution 2.09 Å), were downloaded from the RCSB Protein Data Bank.¹ These files were processed in AutoDockTools-1.5.7 to prepare them for docking simulations (Morris et al., 2009). This involved removing water molecules, other chains, and any previously docked ligands. Polar hydrogens were added, non-polar hydrogens were merged, and the energy minimization of protein residues was performed by adding Kollman charges. Any missing atoms were also repaired as necessary (Seniya et al., 2015).

The 2D structure of Shatavarin IV (CID: 441896), an active biomolecule found in Asparagus racemosus, was downloaded from the PubChem database² in SDF format. The 2D structure coordinates were converted into 3D using Avogadro for docking simulation studies against HIV-1 proteins. Tautomeric and stereochemical modifications were made, and energy minimization of Shatavarin IV was performed using the Avogadro tool (Hanwell et al., 2012). The resulting 3D coordinates were saved in PDB format. Furthermore, the Shatavarin IV 3D coordinates file was processed under AutoDock4.2 to correct torsion angles and perform energy minimization by adding Gasteiger charges.

Molecular docking simulations were conducted to explore Shatavarin IV’s inhibitory potential against HIV-1 proteins. These simulations allow for the study of catalytic behaviour due to molecular interactions between the protein and the ligand Shatavarin IV. The HIV replication protein PDB and ligand 3D files were imported into AutoDock v4.2.³ AutoDock generated a set of ten conformation models representing the ten best models for predicting how Shatavarin IV interacts with HIV-1 Integrase, HIV-1 Protease, and HIV-1 Reverse Transcriptase. Using a Lamarckian genetic algorithm, AutoDock generated various energy values, i.e., binding energy, ligand efficiency, inhibition constant, intermolecular energy, Van der Waals, electrostatic, and total internal energy, used to analyse the relative strengths of the contacts. Binding energy, ligand efficiency, and inhibition constant are crucial indicators of the overall strength of a projected interaction assessed by AutoDock. A mass-centred grid box with a spacing of 0.375 Å was generated using the AutoGrid program and centred on active site pocket residues available in the binding pocket of HIV-1 proteins. These active site residues were identified by observing the bound proteins downloaded from the RCSB Protein database using BIOVIA Discovery Studio Visualizer v21.1.0.20298 software.⁴ These residues were later employed as the ‘flexible’ residues of the macromolecules (i.e., HIV-1 proteins) in the AutoDock experiment.

2D and 3D molecular interactions, along with conformational analysis, were conducted using the PMV tool within AutoDockTools v1.5.7 and the BIOVIA Discovery Studio Visualizer v21.1.0.20298 software. The binding energy was utilized to rank the ten conformations obtained in each simulation run. Conformation #1, characterized by the lowest energy, was identified as the one exhibiting the strongest binding, while conformation #10, with the highest energy, was considered to represent the weakest binding. Following established methodologies (Morris et al., 2009), we scrutinized and interpreted the energy data generated by AutoDock. According to AutoDock, the binding energy is the cumulative sum of intermolecular forces acting on the receptor-ligand complex (Equation 1) (Lin et al., 2011). (1) Δ G binding = Δ G v d w + Δ G elec + Δ G Hbond + Δ G desolv + Δ G torsional

The binding energy obtained can be compared to the calculated Gibbs free energy of the naturally occurring Shatavarin IV and HIV-1 protein interactions (Equation 2), as it essentially reflects a calculated Gibbs free energy value. (2) Binding Energy = Δ G R = 1.987 × 10 − 3 kcal / K ∗ mol = RTlnKT = 298.15 K K a = 0.72 m M = Binding Affinity

In the AutoDock program, the inhibition constant ( K i ) can be computed by first determining the binding energy (ΔG) as described earlier. Subsequently, the inhibition constant ( K i , μM) is calculated using the formula K i = exp Δ G R T , where R represents the universal gas constant ( 1.985 x 10 − 3 Kcal molK ) and T denotes the temperature (298.15 K).

To assess intracellular reactive oxygen species (ROS) generation, particularly superoxide (O2•−), confocal imaging with MitoSOX Red molecular probe was employed, following a previously described method (Jadaun et al., 2022). Briefly, TZM-bl cells (1 × 10⁵) were seeded onto glass coverslips in a 6-well plate, infected with HIV-1_VB028, followed by treatment with AQAR, HAAR and Shatavarin IV for 24 h, and then incubated for 30 min in the dark with MitoSOX (10 μM). Subsequently, the cells were fixed with 3.7% paraformaldehyde and examined using a laser-scanning confocal microscope (Leica TCS SP8) after a phosphate-buffered saline (PBS) wash. A positive control comprising 0.1 mM Xanthine +0.01 U Xanthine oxidase, known ROS producers, was included. Using a standard magnification of 63 × 1.4 NA Oil objective, all confocal images were captured.

Intracellular calcium concentrations, both cytosolic and mitosolic, were determined using the cytosolic calcium indicator Fluo 3 AM and the mitochondria-specific calcium dye Rhod 2 AM, respectively. Confocal microscopy was utilized for qualitative identification of intracellular cytosolic and mitosolic calcium. TZM-bl cells (1 × 10⁵) were seeded onto glass coverslips in 6-well plates, infected with HIV-1_VB028, and then treated with extracts and the pure compound Shatavarin IV for 24 h. Following a 30 min incubation in the dark, cells were stained with 5 μM Fluo 3 AM and 5 μM Rhod 2 AM using 0.1% (v/v) Pluronic® F-127 as a dispersant. After washing with PBS, cells were fixed with 3.7% paraformaldehyde in PBS. The coverslips, mounted on slides, were examined using a confocal microscope (Leica TCS SP8).

To evaluate changes in mitochondrial membrane potential indicative of permeability alterations upon HIV-1 infection, we utilized the JC-1 probe, a fluorescent dye commonly used for this purpose (Invitrogen, Carlsbad, United States). The JC-1 probe exists in two forms: a monomeric form emitting green fluorescence and a J-aggregate form emitting red fluorescence. TZM-bl cells (1 × 10⁵) were cultured in six-well plates following HIV-1 infection and treatments with AR extracts and the bioactive molecule Shatavarin IV at their respective EC₈₀ concentrations for 24 h. Subsequently, the cells were exposed to 1 mL of pre-warmed JC-1 staining solution (5 μg/mL) at 37°C for 20 min. After three washes with PBS and fixation with 3.7% paraformaldehyde in PBS, coverslips mounted on slides were analysed using a confocal microscope, as mentioned above.

The Caspase-Glo 3/7 and Caspase-Glo 9 Assay kits (Promega, WI, United States) were used to detect cellular apoptosis. These assays measure the enzymatic activity of caspases, essential regulators of programmed cell death. Caspase 9 initiates the intrinsic apoptotic pathway, while Caspase 3 and Caspase 7 execute apoptosis. The assays employ specific substrates linked to fluorophores (DEVD for Caspase 3/7 and LEHD for Caspase 9). Upon caspase activation, these substrates are cleaved, releasing fluorophores and generating a fluorescent signal. The increase in fluorescence intensity indicates caspase activity, providing a quantitative measure of apoptosis. To assess apoptosis in HIV-1 infected cells treated with AR extracts and Shatavarin IV, Caspase 3/7 and Caspase 9 activities were measured following the manufacturer’s instructions. The Caspase-Glo reagent, containing substrate and assay buffer, was added directly to the cells. After incubation, the signal was detected using a luminometer, allowing for the quantification of apoptosis in a 96-well culture plate format. Cells treated with Camptothecin (15 μm) were taken as a positive control in the experiment.

The collected data were analysed using GraphPad Prism 9.0 (GraphPad Software, United States) and MS Excel 2021. Each experiment was performed with a minimum of three replicates, and the mean ± standard deviation (SD) was calculated accordingly. Statistical significance was determined using a one-way ANOVA using GraphPad Prism version 9.0, comparing both mock-infected and infected experimental groups. Significance was considered at p < 0.05. In the results of all in vitro experiments, asterisks denote the level of significance, with * representing p < 0.05, ** p < 0.01, or *** p < 0.001.

The cytotoxicity of HAAR and AQAR extracts was initially assessed on TZM-bl cell lines and PBMCs using the MTT quantitative colorimetric assay. Dose-dependent kinetics of the extracts (0.0078–1.00 mg/mL) were evaluated by plotting concentrations against the percentage of cell viability. AQAR was found to be less-toxic (<50%) to both cell lines at concentrations up to 1.0 mg/mL, whereas HAAR exhibited tolerability up to 0.50 mg/mL (Figures 1A,B). The CC₅₀ values for AQAR and HAAR extracts on TZM-bl cells were determined as 1.51 mg/mL and 0.50 mg/mL, respectively. A similar trend was observed in PBMCs, with CC₅₀ values of 0.86 mg/mL and 0.57 mg/mL for the aqueous and hydroalcoholic extracts of AR, respectively (Figure 1C). Additionally, the results of the MTT assays were corroborated by the ATPlite assay in TZM-bl cells (Figures 1D,E), with CC₅₀ values of 1.67 mg/mL and 0.98 mg/mL obtained for AQAR and HAAR, respectively (Figure 1F). Concentrations below the determined cytotoxic levels, as indicated by the CC₅₀ values, were selected for anti-HIV-1 screening.

Cell-associated assays were employed to assess the anti-HIV-1 activity of AQAR and HAAR in the TZM-bl cell line, followed by validation in PBMCs. In these assays, azidothymidine (AZT: 0.49 μM) served as the positive control (data not shown). The luciferase gene assay was utilized in TZM-bl cells to evaluate the antiviral activity of AQAR and HAAR extracts against HIV-1 infection. Separate infections with two distinct HIV-1 subtypes (HIV-1_VB028, subtype C, and HIV-1_UG070, subtype D) were conducted in TZM-bl cells before treatment with various doses of extracts. After 48 h of incubation, dose-dependent inhibition of HIV-1_VB028 and HIV-1_UG070 viral strains was observed with both extracts across different dosages (0.0156–0.50 mg/mL) (Figures 2A,B). The 50% effective concentration or EC₅₀ values for AQAR and HAAR against HIV-1_VB028 were determined to be 0.042 mg/mL, respectively (Figure 2C). For HIV-1_UG070, the EC₅₀ values were found to be 0.069 mg/mL for AQAR and 0.045 mg/mL for HAAR (Figure 2C). These results were further validated in PBMCs using the primary isolate HIV-1_VB028, where a dose-dependent suppression of HIV-1 p24 antigen was evident (Figures 2D,E). In PBMCs, the EC₅₀ values for AQAR and HAAR were found to be 0.072 mg/mL and 0.035 mg/mL, respectively (Figure 2F). Notably, AQAR and HAAR extracts exhibited 80% suppression of HIV-1 infection at non-toxic concentrations, with EC₈₀ doses determined to be 0.199 mg/mL and 0.106 mg/mL, respectively (Figure 2F). The EC₈₀ concentrations of the extracts were subsequently used for enzymatic and confirmatory antiviral efficacy assays against HIV-1.

Additionally, extracellular p24 antigen levels (pg/mL) and HIV-1 viral copy numbers released in the culture supernatants of PBMCs infected with HIV-1 were assessed. At the eleventh-day post-infection (PID), treatment with AQAR and HAAR extracts significantly reduced p24 release by 92.6 to 96% in the supernatant, respectively (Figure 3A). Employing an automated viral load quantitative platform (Abbott m2000 System), the HIV-1 viral copy number was measured, with results depicted as RNA copies/mL (Figure 3B). Treatment with AQAR and HAAR extracts at their respective EC₈₀ doses (0.19 mg/mL and 0.10 mg/mL) resulted in 29,971 copies/mL and 7,304 copies/mL, respectively, compared to the virus control (>10,00,000 copies/mL). The well-established standard drug AZT (0.49 μM) served as the positive control in both experiments.

To elucidate the mechanism of action, an in vitro HIV-1 Integrase assay was conducted to screen both extracts at their respective EC₈₀ concentrations. AQAR exhibited 26.6% inhibition, while HAAR showed 37.3% inhibition compared to the known standard drug, Raltegravir (0.48 μM; SD), which displayed 98.6% inhibition (Figure 4A). HIV-1 integrase facilitates the insertion of viral DNA into the host genome and plays a crucial role in the HIV-1 replication cycle. Notably, neither of the extracts demonstrated significant inhibition of the HIV-1 integrase protein. The kit provided Enzyme Control (EC) and 1.0% Sodium Azide (PC) were employed as the negative and positive controls, respectively.

The ability of AR extracts to suppress HIV-1 protease was evaluated using the in vitro kit-based assay. Results showed that AQAR extract, at a sub cytotoxic dose (0.19 mg/mL), inhibited HIV-1 protease activity by 44.9%, while HAAR (0.10 mg/mL) inhibited its activity by 58.5% (Figure 4B). When compared to Ritonavir (10 μM; SD), a well-known HIV-1 protease inhibitor, which decreased protease activity by 71.0%, both extracts exhibited moderate inhibition of HIV-1 protease. Furthermore, the assay was validated with the kit-provided positive inhibitor (PC), Pepstatin (1 mM), and Enzyme Control (EC).

Reverse transcriptase (RTase) is essential for retroviruses to replicate single-stranded viral RNA. HIV-1 RTase copies the viral RNA into double-stranded DNA, which is then integrated into the host genome. It was found that AQAR extracts inhibited HIV RTase activity by 59.7% at the EC₈₀ dose, whereas HAAR extract inhibited the HIV-1 RTase enzyme by 73.0% (Figure 4C). This was compared to the well-known HIV-1 RTase inhibitor AZT (0.49 μM; SD), which inhibited the HIV-1 reverse transcriptase enzyme by 86.8%. Among the three essential replication enzymes, AR extracts exhibited the maximum reduction in reverse transcriptase enzyme activity.

Molecular docking simulations utilized a key bioactive molecule, Shatavarin IV, from Asparagus Racemosus root extracts, identified through HR-MS using ESI+ method and Orbitrap mass analyzer, along with various HIV-1 proteins. These simulations focused on interactions between Shatavarin IV and HIV-1 proteins due to the inhibitory effects observed in in vitro enzymatic assays. Shatavarin IV was screened based on binding energy (Kcal/mol) and inhibition constant (Ki) value alongside various biomolecular interactions with HIV-1 proteins, including hydrogen bonds and hydrophobic interactions (Table 1). The inhibition constant (Ki) measures the strength of the interaction between a protein and a ligand, with a lower value indicating a lower dissociation and higher inhibition probability (Pandey and Verma, 2022). It is calculated as K i = exp δ G / R ∗ T where δG is the free energy of binding, R is the gas constant (1.987 cal K⁻¹ mol⁻¹), and T is the temperature (298.15 K) (Morris et al., 1998). The molecular interactions demonstrate the tight binding of Shatavarin IV to HIV-1 key proteins present in the active site binding pocket, leading to structural rearrangements that achieve specific conformation and orientation, ultimately resulting in the inhibition of HIV-1 replication; hence, it is considered for further in-depth analysis. The therapeutic potential of Shatavarin IV against cancer and Parkinsonism has been documented (Mitra et al., 2012; Smita et al., 2017), but its antiviral efficacy remains unexplored. This study sheds light on stable molecular interactions between Shatavarin IV and key HIV-1 proteins (Table 1), indicating its potential as a novel antiviral agent.

To verify the bioactivity of Shatavarin IV, the pure compound was initially assessed for cellular viability on TZM-bl cell lines and PBMCs using the MTT quantitative colorimetric assay. Concentration-dependent effects of the extracts (0.039–5.00 mg/mL) were observed, with concentrations plotted against the percentage of cell viability (Figure 7A). The CC₅₀ values were determined to be 0.516 mg/mL for TZM-bl cells and 0.419 mg/mL for PBMCs (Figure 7B). These results were further supported by ATPlite assay results in TZM-bl cells (Figure 7C), with CC₅₀ and CC₂₀ (cytotoxic concentration causes a 20% reduction in cell viability) values for Shatavarin IV determined as 0.405 mg/mL and 0.142 mg/mL, respectively (Figure 7D).

Concentrations ranging below the CC₅₀ value (0.016–0.500 mg/mL) of Shatavarin IV were then assessed to determine its anti-HIV-1 activity using a luciferase gene assay in HIV-1_VB028 infected TZM-bl cells, with further confirmation in PBMCs (Figures 7E,F). Shatavarin IV treatment for HIV-1 suppression yielded EC₅₀ and EC₈₀ values of 0.067 mg/mL and 0.172 mg/mL in TZM-bl cells, respectively, while in PBMCs, the values were 0.035 mg/mL and 0.102 mg/mL (Figure 7G).

Furthermore, treatment with Shatavarin IV at a sub cytotoxic dose (0.10 mg/mL) significantly suppressed p24 antigen (pg/mL) and viral RNA levels by over 98% (98.3–98.8%) in PBMCs supernatant at 11th PID, as confirmed by extracellular HIV-1 p24 antigen (pg/mL) and HIV-1 viral load assays, highlighting its potent role as an inhibitor of HIV-1 (Figures 8A,B). Remarkably, in the HIV-1 viral load assay, Shatavarin IV-treated HIV-1 infected cells showed a count of 11,808 cp/mL, whereas the virus control exhibited >10,00,000 cp/mL (Figure 8B).

To elucidate the probable mechanism underlying the potent anti-HIV-1 activity of Shatavarin IV, we tested its effects on key HIV-1 enzymes in vitro to validate the molecular interaction studies. Shatavarin IV demonstrated significant inhibitory activity against both HIV-1 protease and reverse transcriptase. At a sub-cytotoxic concentration (0.10 mg/mL), Shatavarin IV inhibited HIV-1 protease activity by 62.4%, compared to the standard drug Ritonavir, which reduced protease activity by 77.8% (Figure 8C). Moreover, Shatavarin IV exhibited substantial inhibition of HIV-1 RTase, reducing its activity by 87.6%, compared to AZT, which inhibited RTase by 92.9% (Figure 8D). These results align with molecular interaction studies, where Shatavarin IV displayed strong binding affinity to the active sites of both HIV-1 PR and RTase, suggesting that its inhibitory effects arise from effective molecular interactions with these enzymes, potentially disrupting their catalytic functions. This mechanistic insight underscores Shatavarin IV’s promising role as a bioactive compound in inhibiting HIV-1 replication.

The HIV-1 infection induces oxidative stress and mitochondrial dysfunction by elevating reactive oxygen species (ROS) levels (Ivanov et al., 2016). Dysregulation of mitochondrial calcium exacerbates ROS production and mitochondrial membrane potential alterations (Foo et al., 2022). Using the MitoSOX fluorescent probe, we assessed ROS levels in HIV-1 infected TZM-bl cells to evaluate the antioxidant activity of Asparagus Racemosus extracts and its key bioactive molecule, Shatavarin IV. Confocal microscopy revealed higher mitochondrial ROS content in HIV-1 infected cells compared to mock cells. Meanwhile, treatment with AR extracts and Shatavarin IV reduced fluorescence intensity in HIV-1 infected cells (Figure 9A; Supplementary Figure S2).

Changes in permeability result in a reduction in the potential of the mitochondrial membrane (Ψm), which can be assessed using the JC-1 dye molecular probe (Sivandzade et al., 2019). We evaluated Ψm changes in mock and HIV-1 infected cells in the presence or absence of AR extracts and Shatavarin IV treatment. Confocal microscopy revealed increased fluorescence intensity in HIV-1 infected cells, indicative of Ψm loss. However, treatment with AR extracts and Shatavarin IV restored Ψm levels similar to mock control cells (Figure 9B; Supplementary Figure S3).

Calcium homeostasis plays a critical role in regulating apoptotic machinery. Consequently, calcium accumulation in the mitochondria leads to the opening of the membrane permeability transition pore, resulting in a decrease in the potential of the mitochondrial membrane (Giorgi et al., 2012). To evaluate the distribution of cytosolic and mitosolic calcium in HIV-1 infected TZM-bl cells, we utilized Fluo 3 AM and Rhod 2 AM fluorescent probes. After 24 h of exposure, HIV-1 infected cells exhibited increased fluorescence intensity of both Fluo 3 AM and Rhod 2 AM, indicating elevated levels of cytosolic and mitochondrial calcium, respectively. However, treatment with AR extracts and Shatavarin IV reduced both cytosolic and mitosolic calcium levels, suggesting the restoration of mitochondrial membrane potential (Figure 10A; Supplementary Figure S4).

Caspases, members of the cysteine protease family, play a pivotal role in programmed cell death, triggered notably by HIV-1 envelope proteins (Cicala et al., 2000). Hence, we investigated the activation of caspase-3/7 and caspase-9 to assess the impact of AR extracts and Shatavarin IV in HIV-1 infection. Caspase 3/7 and Caspase 9 assays detect the enzymatic activity of these caspases, where Caspase 9 serves as an initiator caspase in the intrinsic apoptotic pathway, Caspase 3 and Caspase 7 act as effector caspases in the execution phase of apoptosis. We observed an 86% increase in Caspase-3/7 activity in HIV-1 infected cells compared to mock-infected cells (Figure 10B). However, treatment with AQAR and HAAR extracts significantly inhibited this elevated activation by 33.3 and 34.7%, respectively, compared to the virus control, while Shatavarin IV treatment exhibited a 52.9% inhibition in Caspase 3/7 activity in HIV-1 infected cells (Figure 10B). Similarly, Caspase 9 activity increased by over 92% in HIV-1 infected cells compared to mock infection (Figure 10C). However, treatment with AQAR and HAAR extracts, as well as Shatavarin IV, reduced this activation by 87, 88.7, and 86.5%, respectively, compared to the virus control (Figure 10C). Thus, extracts of Asparagus Racemosus and its key bioactive molecule Shatavarin IV mitigated caspase activation in HIV-1 infection. Overall, these results underscore the antiviral potential of AR and Shatavarin IV and their capability to alleviate ROS-mediated mitochondrial dysfunction.