Structural Features Underlying the Mismatch Between Catalytic and Cytostatic Properties in L-Asparaginase from Rhodospirillum rubrum

Abstract

The underlying structural features of the mismatch between catalytic and cytostatic properties in L-asparaginase from Rhodospirillum rubrum (RrA) and three of its mutants were investigated. The rationale for selecting the specific mutations (RrA_{A64V, E67K}; RrA_{R118H, G120R}; RrA_{E149R, V150P, F151T}) is to elucidate the role of inter-subunit interaction in RrA and its impact on catalytic efficiency and stability. Bioinformatic modeling revealed a predominantly negative surface charge on RrA with limited positive charge clusters in the vicinity of the interface region. Thus, some negatively charged groups were replaced with positively charged ones to enhance the electrostatic interactions and stabilize the enzyme quaternary structure. RrA_{A64V, E67K} and RrA_{R118H, G120R} additionally contained an N-terminal 17-amino acid capsid peptide derived from the bacteriophage T7 (MASMTGGQQMGRGSSRQ), which could potentially affect the conformational stability of theenzymes. Circular dichroism (CD) spectroscopy was applied to the kinetic parameters analysis of Asn hydrolysis and showed that native RrA displayed a V_max of 30 U/mg and a K_M of 4.5 ± 0.5 mM. RrA_{E149R, V150P, and F151T} exhibited a substantially increased V_max of 57 U/mg. The catalytic efficiency of V_max/K_M also improved compared to the native enzyme: the V_max/K_M increased from approximately 7 U/mg × mM⁻¹ (for the native enzyme) to 9 U/mg × mM⁻¹ for Mut3. Other mutants exhibited less pronounced changes. Thermo-denaturation studies allowed us to determine the phase transition parameters of the RrA variants in comparison with commercial reference sample EcA. RrA_{A64V, E67K} and RrA_{R118H, G120R} exhibited the most favorable phase transition parameters, with melting temperatures (T_m) of 60.3 °C and 59.4 °C, respectively, exceeding that of the wild-type RrA (54.6 °C) and RrA_{E149R, V150P, F151T} (52 °C). The EcA demonstrated a slightly superior thermal stability, with a T_m of 62 °C. The mutations showed a significant effect on protein stability during trypsinolysis. Therefore, RrA_{E149R, V150P, F151T} showed higher resistance (45% activity remaining after 30 min of trypsin exposure) compared to the native RrA retained 20% activity. EcA preparations exhibited lower stability to trypsinolysis (losing over 90% activity in 15 min). The cytostatic effects were evaluated using MTT assays against K562 (leukemic) and A549 (lung carcinoma) cell lines. The MTT assays with K562 cells revealed that RrA_{E149R, V150P, F151T} (IC₅₀ of 10 U/mL) and RrA_{R118H, G120R} (IC₅₀ of 11.5 U/mL) exhibited superior antiproliferative activity compared to native enzymes RrA (IC₅₀ of 15 U/mL) and EcA (24 U/mL). RrA_{E149R, V150P, F151T} showed the most significant improvement in cytostatic activity. The results obtained indicate that the substitutions in RrA_{E149R, V150P, F151T} resulted in the improvement of the enzyme biocatalytic properties and an increase in the resistance to aggregation and trypsinolysis. This highlights the role of electrostatic interactions in stabilizing the oligomeric structure of the enzyme, which eventually translates into an improvement in cytostatic efficiency and antiproliferative forces.

1. Introduction

L-asparaginase, an enzyme catalyzing the hydrolysis of L-asparagine to L-aspartic acid and ammonia, is a valuable component in the treatment of acute lymphoblastic leukemia (ALL) and certain lymphomas [1,2,3,4,5,6]. Bacterial L-asparaginases from Escherichia coli (EcA) and Erwinia chrysanthemi (ErA) are currently employed in combination chemotherapy, exploiting the dependence of rapidly proliferating tumor cells on exogenous asparagine [7]. This dependence is particularly pronounced in leukemic cells, which exhibit significantly lower asparagine synthetase activity compared to normal tissues [8,9,10,11,12,13,14,15,16,17,18]. L-asparaginase treatment leads to cell cycle arrest in the G1 phase and apoptosis. Bacterial L-asparaginases are broadly classified into two types [19,20,21]:

• Type I, constitutively expressed cytoplasmic enzymes with high K_M (~1–8 mM) for L-asparagine, generally lacking significant anti-tumor activity (e.g., Bacillus subtilis, Methanococcus jannaschii, and Pyrococcus horikoshii);
• Type II, periplasmic enzymes with low K_M (~10–20 µM) for L-asparagine, demonstrating antiproliferative activity and including those used clinically (EcA and ErA), along with others such as Yersinia pseudotuberculosis, Helicobacter pylori, and Wolinella succinogenes.

L-asparaginase from Rhodospirillum rubrum (RrA) is noteworthy as a Type I cytoplasmic enzyme that, unlike most others in its class, exhibits demonstrable antiproliferative activity. The crystal structure of the RrA protein has recently been elucidated, as have the structures of several mutant forms, which exhibit topologically unique tetramers [22]. However, a critical consideration in the clinical application of L-asparaginases is the avoidance of significant L-glutaminase activity, which can contribute to toxicity [23,24,25,26]. While EcA and ErA are currently the only bacterial L-asparaginases approved for medical use, their L-glutaminase activity (5–10% of L-asparaginase activity) and relatively high K_M for L-glutamine (1–3 mM) necessitate careful monitoring for adverse effects.

RrA presents a compelling alternative due to its significantly lower L-glutaminase activity (<0.01% of L-asparaginase activity) and its low homology to EcA and ErA [3,27,28,29,30,31]. This low homology, coupled with its unique structural features—including a shorter amino acid sequence (172 residues per monomer compared to 330 in EcA and ErA)—makes RrA a potential candidate for treating patients who develop hypersensitivity to clinically used asparaginases. Further highlighting its potential, RrA demonstrates cytotoxic activity against several human leukemia (K562), prostate cancer (DU145), and breast cancer (MDA-MB-231, MCF7) cell lines, surpassing the cytotoxic effects of ErA while exhibiting lower overall toxicity compared to EcA [30]. The present study aims to characterize the physicochemical properties, biocatalytic characteristics, and cytostatic efficacy of RrA and three of its mutants in comparison with established commercial L-asparaginase preparations (EcA). This detailed characterization will contribute significantly to the evaluation of RrA’s potential as a novel therapeutic agent for leukemia.

The oligomeric state is a critical determinant of both biocatalytic properties and stability in asparaginases. Most bacterial L-asparaginases exist as tetramers with four active sites, each formed at the interface of two subunits [31,32,33,34,35]. While the dimeric form retains some L-asparaginase activity, it is significantly less active than the tetramer [36]. Stabilizing the tetrameric structure of RrA through inter-subunit crosslinking with bifunctional agents like spermine and spermidine significantly enhanced both catalytic and antitumor activity [37]. Bioinformatic studies have revealed a predominantly negative surface charge on RrA with limited positive charge clusters [38].

So, the rationale for selecting the specific mutations RrAA64V, E67K; RrAR118H, G120R; RrAE149R, V150P, F151T stemmed from a desire to elucidate the role of inter-subunit interactions in RrA and their impact on catalytic efficiency and stability. In the case of the substitutions studied, it is expected that the rigidity and stability of the protein globule packing will increase by means of additional salt linkage and the saturation of the protein molecule’s nonpolar contacts. In the presented work, we aimed to enhance electrostatic interactions by reducing negative charge density in the region of inter-subunit interactions. Mutants RrA_{A64V, E67K}, RrA_{R118H, G120R}, and RrA_{E149R, V150P, F151T} were studied in which reduction of negative charges near areas of contact of both subunits is implemented, which may lead to reduced repulsion monomers generally negatively charged and increase stability of the whole macromolecule.

RrA and its mutant variants were studied, as well as the influence of the N-terminal peptide: the 17-amino acid peptide sequence, MASMTGGQQMGRGSSRQ, is a capsid peptide derived from the bacteriophage T7 and has been engineered to enhance the thermostability of RrA. T7 tag inclusion was based on established literature precedent for its utility in protein engineering applications, including labeling, purification, and characterization [32,39]. The T7 tag is relatively hydrophilic and could indirectly enhance thermostability by preventing aggregation. So, the presence of the T7 tag results in an enhancement of the thermal stability and resistance to denaturation of several mutant forms of the RrA protein in the presence of urea [39]. Consequently, we propose that the N17 terminal amino acid sequence of the capsid protein of bacteriophage T7 tag may also contribute to enhanced thermal stability in other RrA mutant forms.

Here, we investigated the correlation between the physicochemical kinetic properties of RrA and its mutants with the cytotoxicity potential. We evaluated alterations in thermostability, determining the impact of mutations on the enzyme’s resistance to thermal denaturation and its potential for enhanced stability in vivo. Furthermore, we assessed the impact of mutations on V_max, with substrate specificity (K_M) providing crucial insights into the catalytic efficiency of each mutant. This investigation contributes to the rational engineering of improved enzyme-based therapeutics.

2. Results and Discussion

2.1. FTIR Spectroscopy for Characterization of Native and Mutant RrA L-Asparaginases and Comparison with Pharmaceutical (EcA) Preparations

FTIR spectroscopy was used to characterize native and mutant forms of RrA L-asparaginase (

Table 1. The characteristics of mutant forms of L-Asparaginase RrA from Rhodospirillum rubrum.

Designation	Substitutions in DNA	Substitutions in Protein
Mut1 *	C191T, G199A	A64V, E67K
Mut2 *	G353A, G354C, G358A	R118H, G120R
Mut3	G445A, A446G, G448C, T449C, T450C, T451A, T452C	E149R, V150P, F151T

* The Mut1 and Mut2 mutants contain the N-fragment MASMTGGQQMGRGSSRQ.

) and compare them to commercially available asparaginases (EcA).

Figure 1a displays the FTIR spectra within the 1800–900 cm⁻¹ range. The characteristic Amide I band (1700–1590 cm⁻¹) and Amide II band (1580–1500 cm⁻¹) protein major bands have been observed for the presented proteins [40]. The FTIR spectral analysis of all asparaginases reveals a remarkable degree of similarity, with a notable proximity in the positions of the centroids of the peak maxima for the Amide I and Amide II bands. These centroids fall within the ranges of 1640–1647 cm⁻¹ for Amide I and 1540–1553 cm⁻¹, which is consistent with expectations for proteins belonging to similar classes. The native RrA exhibits Amide I and Amide II mass centers at 1643.2 cm⁻¹ and 1549 cm⁻¹, respectively, corresponding to pronounced β-sheet impact. The RrA_{R118H, G120R} and RrA_{A64V, E67K} show slight shifts to lower wavenumbers in the Amide I band (1641.3 cm⁻¹ and 1641 cm⁻¹, respectively), suggesting some alterations in secondary structure due to the mutations. These FTIR data highlight structural variations in secondary structure between native and mutant RrA L-asparaginases and commercially available EcA preparations that may contribute to differences in enzyme activity and stability among these different L-asparaginase preparations.

Figure 1b,c present the Gaussian deconvolution of the Amide I band to determine secondary structure content for RrA and its mutants in comparison with EcA preparations. Figure 1d shows the secondary structure composition.

Compared to native RrA, the mutant forms (RrA_{A64V, E67K} and RrA_{R118H, G120R}) show differences in their secondary structure. The mutants show a decrease in α-helices and a substantial increase in β-sheet content compared to native enzymes, especially RrAR118H and G120R (28.12% vs. 18% for native RrA) (Figure 1d). An increase in b-sheet was also observed for RrA_{R118H, G120R}, which is probably due to the additional antiparallel b-sheet formed in the intersubunit region.

There is a clear discrepancy between RrA and commercial EcA. EcA-Medac has a higher α-helix content (38.1%) and lower random coil content (21.8%) than RrA. EcA-Veropharm shows the average values of the α-helix content (27.5%). A similar percentage of α-helices and β-structural elements was obtained by analyzing the CD spectra for RrA with the proportion of α-helices at 36 ± 2%, β-sheet 14 ± 4%, β-turns 16 ± 2%, and random coil 30 ± 3% [41]. For the commercial drug EcAII (Kidrolase), the proportion of α-helices is 32.2% and18.3% for the β-sheet [42]. This allows us to come to a conclusion about the effect of mutations on the structure and, consequently, the function of the enzyme.

2.2. Determination of the Catalytic Parameters of L-Asparaginase RrA and Its Mutant Forms

2.2.1. 3D Structures of RrA and Mutant Forms to Visualize How Amino Acid Substitutions Could Affect the Conformation Stability of L-Asparaginase

The crystal structure of the RrA protein has recently been elucidated, as have the structures of several mutant forms, which exhibit topologically unique tetramers [22]. RrA and EcA share similar overall topologies, with a key distinction residing in the central β-sheet. Notably, RrA exhibits a more ordered active site loop compared to EcA, and these active site loops play a crucial role in stabilizing the tetrameric structure. RrA could form homotetramers at concentrations as low as 25 nM. Mutations of key residues within the active site, specifically Lys19, Tyr21, and Lys158, have been shown to impact catalysis. Furthermore, the crystallization of these mutants in the absence or presence of L-Asp results in distinct conformational states [22].

The rationale for selecting the specific mutations RrA_{A64V, E67K}; RrA_{R118H, G120R}; RrA_{E149R, V150P, F151T} stemmed from a desire to elucidate the role of inter-subunit interactions in RrA and their impact on catalytic efficiency and stability.

In the presented work, mutants RrA_{A64V, E67K}, RrA_{R118H, G120R}, and RrA_{E149R, V150P, F151T} were studied in which a reduction of negative charges near areas of contact of both subunits was implemented, which may have led to a reduction in generally negatively charged repulsion monomers and an increase in stability of the enzyme structure. On the other hand, the rigidity and stability of the protein globule packing can be achieved by means of additional salt linkage and the saturation of the protein molecule nonpolar contacts. The location of the substitutions within the RrA structure is illustrated in Figure 2.

2.2.2. Kinetics Curves

Circular dichroism (CD) spectroscopy offers a unique approach to determining L-asparaginase catalytic parameters by monitoring the enzymatic hydrolysis of L-asparagine. By establishing a calibration curve relating the CD signal to substrate concentration (the difference in the ellipticity of L-asparagine and L-aspartic acids with a concentration at λ = 210 nm Δε = 1.66 mdeg/mM), the reaction kinetics can be determined, enabling the calculation of kinetic parameters V_max and K_M. This provides a continuous, real-time measurement of enzymatic activity, offering advantages over endpoint assays, such as the direct Nesslerization method [43]. Thus, CD spectroscopy offered distinct advantages for assessment of L-asparaginase activity due to the absence of chromogenic substrate, particularly in its ability to provide continuous, reagent-free monitoring under a wider range of conditions, making it well-suited for high-throughput screening and characterization.

The main limitation of the CD spectroscopy method is the requirement for optical transparency of the sample (it is not possible to measure activity in blood serum or whole blood).

Figure 3a exemplifies the kinetic curves of Asn hydrolysis catalyzed by native RrA L-asparaginase and three mutant variants, RrA_{A64V, E67K}, RrA_{R118H, G120R}, and RrA_{E149R, V150P, F151T}, obtained using the CD spectroscopy method. These curves demonstrate the time-dependent decrease in the CD signal reflecting the conversion of L-asparagine to L-aspartic acid. The Michaelis–Menten kinetics scheme was applied to the data to determine the kinetic parameters. Analysis was performed using both nonlinear regression (Figure 3b) and three linearization methods: Lineweaver–Burk (Figure 3c), Hanes–Woolf (Figure 3d), and Eadie–Hofstee (Figure 3e).

Table 2. Kinetic parameters K_M and V_max of Asn hydrolysis by L-asparaginase RrA native and mutant forms, determined using nonlinear regression and linear approximation in various coordinates. The experimental conditions are similar to those indicated in the caption of Figure 2.

Michaelis–Menten nonlinear regression analysis
Enzyme	Native RrA	RrAA64V, E67K (RrA Mut1)	RrAR118H, G120R (RrA Mut2)	RrAE149R, V150P, F151T (RrA Mut3)
Vmax, U/mg	29.6 ± 1.4	31.6 ± 0.7	34.9 ± 0.4	57.2 ± 2.4
KM, mM	4.5 ± 0.5	5.0 ± 0.3	6.7 ± 0.4	6.5 ± 0.7
R-Square	0.9940	0.9979	0.9998	0.9970
Lineweaver–Burk plot
Enzyme	Native RrA	RrAA64V, E67K (RrA Mut1)	RrAR118H, G120R (RrA Mut2)	RrAE149R, V150P, F151T (RrA Mut3)
Vmax, U/mg	29.7 ± 1.8	31.7 ± 0.8	34.7 ± 0.4	56.4 ± 2.2
KM, mM	4.3 ± 0.7	5 ± 0.4	6.5 ± 0.4	6.1 ± 0.7
R-Square	0.9428	0.982	0.9832	0.9715
Hanes–Woolf plot
Enzyme	Native RrA	RrAA64V, E67K (RrA Mut1)	RrAR118H, G120R (RrA Mut2)	RrAE149R, V150P, F151T (RrA Mut3)
Vmax, U/mg	29.7 ± 1.8	31.7 ± 0.8	34.7 ± 0.4	56.4 ± 2.2
KM, mM	4.3 ± 0.2	5 ± 0.1	6.5 ± 0.1	6.1 ± 0.1
R-Square	0.9793	0.9962	0.9991	0.9912
Eadie–Hofstee diagram
Enzyme	Native RrA	RrAA64V, E67K (RrA Mut1)	RrAR118H, G120R (RrA Mut2)	RrAE149R, V150P, F151T (RrA Mut3)
Vmax, U/mg	30.9 ± 2.3	31.8 ± 0.8	33.9 ± 0.4	57.6 ± 5.6
KM, mM	5.1 ± 0.5	5.1 ± 0.2	5.7 ± 0.3	6.9 ± 1.2
R-Square	0.9454	0.9932	0.9843	0.8436

presents the kinetic parameters (K_M and V_max) for Asn hydrolysis by L-asparaginase RrA native and mutant forms.

A more detailed, nonlinear Michaelis–Menten regression analysis (Table 2) revealed a V_max of 29.6 ± 1.4 U/mg for the native RrA enzyme and K_M of 4.5 ± 0.5 mM. The mutant RrA_{E149R, V150P, F151T} exhibited a significantly increased V_max of 57.2 ± 2.4 U/mg, approximately double that of the native enzyme, indicating a substantial enhancement in catalytic activity. This increase in V_max was accompanied by a slightly elevated K_M of 6.5 ± 0.7 mM, suggesting a modest reduction in substrate affinity. Mutants RrA_{A64V, E67K} and RrA_{R118H, G120R} showed less dramatic changes in V_max (31.6 ± 0.7 U/mg and 34.9 ± 0.4 U/mg respectively) with K_M values of 5.0 ± 0.3 mM and 6.7 ± 0.4 mM respectively, indicating smaller alterations in both catalytic efficiency and substrate affinity compared to Mut3. Amino acid substitutions near position 150 have a more pronounced effect on the catalytic parameters. All nonlinear regression analyses displayed high R-squared values (≥0.9970), confirming their excellent fits to the Michaelis–Menten model.

The Lineweaver–Burk, Hanes–Woolf, and Eadie–Hofstee linearization methods (Table 2) yielded comparable V_max values to the nonlinear regression. However, the K_M values displayed minor variations across these methods, underscoring the inherent limitations and potential biases of linearization approaches. In particular, the Eadie–Hofstee plot for Mut3 showed a substantially lower R-squared value (0.8436), indicating a poor fit, which further supports the superior reliability of nonlinear regression for precise kinetic parameter determination. This consistency was observed between the nonlinear regression results and the other methods. However, the native enzyme and mutants 1 and 2 highlight the overall reliability of the CD-spectroscopy-based approach.

While the mutants do exhibit higher K_M values compared to the wild-type enzyme, it is crucial to consider the physiological concentration of asparagine. In the bloodstream, asparagine concentrations are typically in the micromolar range (40–80 µM) [44]. Given that the K_M values for all of the enzymes studied, including native RrA and its mutants, are higher than this physiological concentration, the enzymes in the bloodstream work far from the saturation mode of the substrate. In this case, catalytic efficiency should be considered an analytically significant parameter (V_max/K_M).

The catalytic efficiency (V_max/K_M) increased from approximately 7 U/mg × mM⁻¹ (for the native enzyme) to 9 U/mg × mM⁻¹ for Mut3. Conversely, Mut1 and Mut2 exhibited a slight decrease in catalytic efficiency to 5–6 U/mg × mM⁻¹. The increased V_max/K_M suggests that Mut3 can process asparagine at a much faster rate, potentially leading to a more rapid and effective therapeutic response.

2.3. The Thermograms of L-Asparaginases and Parameters of Its Thermodenaturation

The thermal stability of native RrA L-asparaginase and its mutant forms, alongside commercial EcA preparations, was assessed using CD spectroscopy. This method effectively monitors protein denaturation by tracking changes in ellipticity as a function of temperature. Protein unfolding, often a cooperative process, can be monitored through CD spectroscopy.

To investigate the structural changes during denaturation, CD spectra were recorded at various temperatures during the thermal scans (Figure 4a). The deconvolution of these spectra (Figure 4b) confirmed the relatively minor changes in secondary structure content observed upon heating native RrA, supporting the initial observation that the overall secondary structure is largely preserved during thermal denaturation.

Thermal denaturation profiles (thermograms) were generated by heating samples from 20 °C to 100 °C at a constant rate of 1 °C/min, followed by cooling at the same rate (Figure 5a). The thermograms of native RrA and its mutants revealed a relatively small change in ellipticity during phase transition (Δθ = 1–8 mdeg) upon heating, indicating the retention of a significant proportion of α-helices and β-sheets.

In the case of pharmaceutical preparations based on EcA, there was a dramatic change in ellipticity within a narrow temperature range close to 60 °C. For EcA Medac, this change amounted to approximately 24 millidegrees (75% change in signal from initial), while for EcA Veropharm, it is approximately 4 millidegrees (35% change in signal from initial). In the case of EcA drugs, an abrupt phase transition occurs. In the case of native RrA and Mut3, the phase transition was smooth, which makes it possible to restore the initial activity during cooling. For RrA-Mut1 and RrA-Mut2, a sharp phase transition occurred with a loss in activity.

The melting temperature (T_m), defined as the temperature at which 50% of the protein is denatured, was estimated by analyzing the CD signal at 220–222 nm, a wavelength range characteristic of α-helices. The T_m values were 54.6 °C for RrA, 52 °C for RrA-Mut3, 59.4 °C for RrA-Mut2, and 60.3 °C for RrA-Mut1, indicating that the last two possessed the highest thermal stability. The temperature of the RrA phase transitions was close to the temperature optimum but at the same time, slightly higher by about 5 °C. Conversely, both EcA enzymes maintained their stability up to 60 °C, with a melting point estimated at 62 °C, showcasing their remarkable thermal stability. Nonetheless, under physiological conditions, RrA exhibited sufficient performance as well.

The cooling experiments demonstrated partially reversible denaturation for RrA variants (Figure 5a,b). For the native RrA and Mut1, the Far-UV CD spectra demonstrated incomplete recovery of the secondary structure after cooling, indicative of aggregation. An analysis of the native RrA CD spectra across a temperature gradient (Figure 4b) revealed a decrease in the α-helical content and an increase in the intermolecular antiparallel β-sheet structure, characteristic of protein aggregation. This observation aligns with the partial recovery (40–50%) observed in the cooling curve of the native RrA thermogram.

In contrast, Mut2 and Mut3 exhibited a higher recovery of secondary structure (70–80%) based on the Far-UV CD spectra before melting and after cooling, as these mutants demonstrated the extent of reversibility (Figure 5b). The reversibility of denaturation observed in Mutants 2 and 3 strongly suggests the absence of aggregation and points towards a subtly altered denaturation mechanism. Remarkably, Mutant 3 retained a significant proportion of its alpha-helical structure even at 100 °C, exhibiting a distinctly alpha-structured spectrum and demonstrating a high degree of secondary structure recovery. Mutant 2 also displayed substantial resilience, recovering over 80% of its secondary structure upon cooling. These findings underscore the stabilizing effect of the cation substitutions at the subunit interface, preventing irreversible aggregation and promoting refolding to the native conformation.

The thermodenaturation process of asparaginases was modeled as an equilibrium between native (N) and denatured (D) states: N <--> D, with an equilibrium constant of K = [N]/[D]. The analysis started with experimental thermograms, which are fitted to an asymmetric sigmoidal function (Figure 5a). Then, the Gibbs free energy change (ΔG) was calculated using the following equation: ΔG = −RT ln K. The enthalpy change (ΔH) is then determined using the van ’t Hoff isobar equation: dlnK/dT = ∆H/(RT²). Finally, the entropy change (ΔS) was calculated using the equation ΔS = (ΔH − ΔG)/T.

Figure 6 presents the thermodynamic analysis of L-asparaginase denaturation. Panel (a) shows the dependence of the natural logarithm on the equilibrium constant (ln K) for the native-to-denatured transition on the temperature (T) following the van ’t Hoff equation. The slope of this plot yielded the enthalpy change (ΔH) of denaturation. Panel (b) displays the enthalpy change (ΔH) as a function of temperature, illustrating its temperature dependence. Panel (c) presents the entropy change (ΔS) as a function of temperature, reflecting the change in disorder upon denaturation.

Thermodynamic parameters presented in

Table 3. Thermodynamic parameters of L-asparaginase thermodenaturation at 37 °C. PBS (pH 7.4, 0.01 M).

Parameter	Native RrA	RrAA64V, E67K (RrA Mut1) *	RrAR118H, G120R (RrA Mut2) *	RrAE149R, V150P, F151T (RrA Mut3)	EcA Medac	EcA Veropharm
ln K	−1.85	−3.73	−1.89	−2.08	−6.04	−5.37
ΔH, kJ/mol	91.1	132.2	97.8	60.2	25.1	26.6
ΔS, J/mol/K	−15.1	−30.6	−15.4	−17.1	−50.1	−44.6

* The Mut1 and Mut2 mutants contain the N-fragment MASMTGGQQMGRGSSRQ.

reveal interesting differences in the stability of native RrA and its mutants at 37 °C. The RrA-Mut1 displays a significantly higher enthalpy of denaturation (ΔH) compared to native RrA, indicating stronger intramolecular interactions within the mutant enzyme. This increase in ΔH is, however, offset by a larger decrease in entropy (ΔS), suggesting a more ordered denatured state. Consequently, the equilibrium constant for denaturation (K) is only slightly lower for Mut1 than for the native enzyme, implying a comparable overall stability at 37 °C. This suggests that the presence of the N-terminal fragment in Mut1 significantly enhances thermostability.

RrA-Mut2 and RrA-Mut3, on the other hand, exhibited a ΔH comparable to native RrA but a smaller decrease in ΔS, resulting in a marginally lower K value and potentially slightly higher stability.

Comparing the RrA variants to the commercially available E. coli asparaginases (EcA Medac and EcA Veropharm) further highlights the differences in their thermodynamic properties. Both EcA variants exhibited considerably lower ΔH and more negative ΔS values compared to native RrA and its mutants. This translates to significantly higher K values for the EcA enzymes, indicating lower stability at 37 °C. These findings underscore the importance of understanding the thermodynamic basis of enzyme stability, which can be influenced by seemingly subtle changes in amino acid sequence, ultimately impacting their suitability for therapeutic applications. The presence of the N-terminal fragment in Mut1 and Mut2 does not appear to drastically alter their thermodynamic profiles compared to native RrA, suggesting that the observed changes are primarily attributable to the specific point mutations introduced.

The literature data for ovalbumin variants showed a correlation between higher ΔH values and higher denaturation temperatures, indicating greater thermostability [40]. The native ovalbumin had a ΔH of 514 kJ/mol and a denaturation temperature of 78.3 °C. Modifications like S-ovalbumin formation (thermostable albumin variant) increase both ΔH (602 kJ/mol) and denaturation temperature (86.0 °C), enhancing stability. Other modifications, such as methylation and lyophilization, decreased both parameters, leading to lower stability. Thus, comparing the asparaginase data with the ovalbumin data highlights the ΔH values for the asparaginases are generally much lower than those of ovalbumin, suggesting that asparaginases are less thermostable.

Thermodynamic analysis reveals minimal losses in thermostability for the RrA asparaginase mutants, while key improvements are observed in other crucial properties. Mutants 2 and 3 exhibit enhanced reversibility to thermodenaturation, which is vital for protein stability in blood plasma and improved kinetic parameters. These findings, combined with the retained thermostability, indicate the successful development of enhanced asparaginase variants with improved therapeutic potential.

2.4. Resistance of L-Asparaginases to Trypsinolysis

The susceptibility of L-asparaginases to trypsinolysis is a crucial factor determining their stability and efficacy in vivo, as trypsin-like proteases in the bloodstream can degrade the therapeutic enzyme. Moreover, the resistance of the enzyme to trypsinolysis correlates with its accessibility to plasma proteins of the complement system. This study investigated the trypsinolytic stability of native RrA and several mutants, comparing them to commercially available EcA preparations (Medac and Veropharm). We hypothesized that trypsinolysis could proceed via two main pathways: denaturation (a first-order process) and/or aggregation (a second-order or higher-order process). To distinguish between these mechanisms, the kinetic data were analyzed using first-order (ln[A] vs. time) and second-order (1/[A] vs. time) plots. Linearity in a first-order plot would suggest denaturation as the predominant mechanism, while linearity in a second-order plot would indicate aggregation.

Figure 7a shows the kinetic curves of trypsinolysis. As shown in Figure 7b, these curves were linearized in first-order coordinates (without aggregation). This indicates that trypsin’s action deliberately leads to structural degradation of the L-asparaginase molecule rather than promoting intermolecular aggregation. The rate of inactivation varied significantly among the different L-asparaginase variants. Native RrA and its mutant RrA-Mut3 exhibited the highest resistance to trypsinolysis, retaining approximately 21% and 45% of their relative activity, respectively, after 60 min (Figure 7). All other L-asparaginases tested showed a more rapid decline in activity, losing over 90% within 15 min.

Table 4. Constants of L-asparaginase inactivation during trypsinolysis (k_in, min⁻¹). The conditions are similar to those shown in Figure 6.

Enzyme	kin, min−1
RrA native	0.025 ± 0.001
* RrAA64V, E67K	0.195 ± 0.013
* RrAR118H, G120R	0.338 ± 0.012
RrAE149R, V150P, F151T	0.015 ± 0.001
EcA-Medac	0.150 ± 0.010
EcA-Veropharm	0.273 ± 0.021

* The Mut1 and Mut2 mutants contain the N-fragment MASMTGGQQMGRGSSRQ.

presents the inactivation rate constants (k_in) for various L-asparaginases during trypsinolysis.

These k_in constants quantify the rate at which each enzyme loses activity in the presence of trypsin, reflecting their susceptibility to proteolytic degradation. A lower k_in value indicates greater resistance to trypsinolysis. The native RrA displays a relatively low k_in (0.025 ± 0.001 min⁻¹), indicating substantial resistance to trypsin. In contrast, mutants RrA_{A64V, E67K} (0.195 ± 0.013 min⁻¹) and RrA_{R118H, G120R} (0.338 ± 0.012 min⁻¹) showed considerably higher inactivation rates, suggesting an increased susceptibility to trypsin. Interestingly, RrA_{E149R, V150P, F151T} demonstrated a surprisingly high stability (0.015 ± 0.001 min⁻¹), even exceeding that of the native enzyme.

The commercial EcA preparations also exhibited relatively high inactivation rates: EcA-Medac (0.150 ± 0.010 min⁻¹) and EcA-Veropharm (0.273 ± 0.021 min⁻¹), comparable to some of the RrA mutants. This suggests that the structural features making these commercial preparations susceptible to trypsinolysis are distinct from those affecting the RrA mutants and native enzymes.

While the N-terminal fragment appears to enhance the thermostability in Mut1 and Mut2, the observed decrease in trypsin resistance requires further discussion. One possibility is that the N-terminal fragment, while stabilizing the protein core, might also introduce or expose trypsin-sensitive sites (arginine and lysine residues) or alter existing loop conformations, making them more accessible to trypsin. This is particularly relevant considering the presence of arginine in the N-terminal sequence (MASMTGGQQMGRGSRQ), which could serve as trypsin cleavage sites.

Integrating the structural model into our analysis provides further insight. The observed differences in trypsinolysis resistance among the mutants likely arise from the specific location and nature of each mutation. In Mut1 and Mut2, the introduction of lysine and arginine, respectively, could create additional trypsin cleavage sites, resulting in enhanced degradation. The substitutions in RrA_{E149R, V150P, F151T} show increased trypsin resistance by inducing localized conformational changes that hinder trypsin access. While E149R introduces another arginine, its location within the model could be sterically hindered or involved in salt bridges, reducing its accessibility to trypsin. The V150P substitution, with proline’s known rigidity, could restrict loop flexibility and limit trypsin accessibility. F151T could alter local hydrophobicity, potentially disrupting favorable interactions with trypsin.

The data in Figure 7c show a positive correlation between the trypsin inactivation rate constant (k_in) and the melting temperature (T_m) for the L-asparaginases studied (which indicate the effect of protein globule compactness and density on stability to trypsinolysis). Higher T_m (greater thermal stability) unexpectedly correlates with higher k_in (lower trypsin resistance). This suggests that while overall compactness may protect some cleavage sites, it might also create less stable regions and strategies to inadvertently increase T_m and trypsin accessibility.

In conclusion, this trypsinolysis study highlights the significant differences in stability among native RrA, its mutants, and commercial EcA preparations. The enhanced stability and catalytic activity of RrA-Mut3 identify it as a promising candidate for further development as a novel therapeutic agent. The preferential denaturation pathway under the conditions of this experiment supports further investigation into the specific trypsin cleavage sites and their influence on overall protein stability. The contrasting behavior of EcA preparations further emphasizes the importance of considering the influence of specific amino acid sequences and protein structure on the in vivo stability and efficacy of L-asparaginase therapeutics.

2.5. Cytostatic Effect of L-Asparaginases on Cancer Cells

We investigated the cytostatic effects of various L-asparaginases on K562 (leukemic) and A549 (lung cancer) cells using MTT assays (Figure 8). The results demonstrate the potent antiproliferative activity of all the tested L-asparaginases against K562 cells. This is consistent with the known dependence of many leukemic cells on exogenous asparagine for survival; therefore, L-asparaginase’s depletion of asparagine effectively inhibits their growth. However, the data also reveal significant variation in potency among enzymes.

In contrast to the K562 results, Figure 8b demonstrates a significantly weaker cytotoxic effect of all L-asparaginases on the A549 cells. This suggests that A549 cells are less dependent on exogenous asparagine or possess mechanisms to compensate for asparagine depletion. The low sensitivity of the A549 cells highlights a key limitation of L-asparaginase-based therapies: their effectiveness is highly dependent on the specific tumor type and its metabolic characteristics. Solid tumors, unlike many leukemias, often exhibit robust asparagine biosynthesis pathways, rendering them less susceptible to L-asparaginase-induced asparagine deprivation.

Table 5. The IC₅₀ values on K562 cells (concentration of semi-inhibition of cell growth) determined using MTT analysis under the action of L-asparaginases. The RPMI-1640 medium with the addition of 5% fetal bovine serum and 1% sodium pyruvate at 5% CO₂/95% air in a humidified atmosphere at a temperature of 37 °C.

Enzyme	IC50 on K562 Cells, U/mL
RrA native	15 ± 2
RrAA64V, E67K *	25 ± 3
RrAR118H, G120R *	11.5 ± 0.7
RrAE149R, V150P, F151T	10 ± 1
EcA-Veropharm	24 ± 4

* The Mut1 and Mut2 mutants contain the N-fragment MASMTGGQQMGRGSSRQ.

presents the IC₅₀ values (concentrations required to inhibit cell growth by 50%) for the K562 cells. The native RrA displayed an IC₅₀ of 15 ± 2 U/mL. Mutants 2 and 3 RrA_{E149R, V150P, F151T} and RrA_{R118H, G120R} showed the strongest cytostatic effect with an IC₅₀ of 10 ± 1 U/mL and 11.5 ± 0.7 U/mL, indicating enhanced anti-leukemic activity compared to the native enzyme. The RrA_{A64V, E67K} exhibited a weaker effect (IC50 = 25 ± 3 U/mL). The commercial preparation of EcA-Veropharm shows an IC₅₀ of 24 ± 4 U/mL, demonstrating comparable activity to RrA_{A64V, E67K} but significantly less potency than the most effective mutants. The N-terminal region in mutant forms of RrA does not significantly affect cytostatic activity. In the context of A549 cell lines, mutant variants exhibit a higher degree of activity compared to native RrA, resulting in a decrease in cell viability ranging from 5% to 15%. Nonetheless, the cytostatic activity in these cells remains inferior to that observed in K562 cell lines.

The difference in the sensitivity (Figure 8a,b) of cancer cell lines to L-asparaginase-induced asparagine depletion is a multifaceted phenomenon that is influenced by a multitude of factors, including the levels of expression of asparagine synthetase (ASNS) [8,9,11,13,15,18,26,45,46,47,48]. An increase in ASNS expression can effectively confer resistance to L-asparaginase, allowing cells to bypass the depletion of extracellular asparagine. Research has demonstrated a correlation between elevated ASNS expression and resistance to L-asparaginase therapy in various types of cancer cells, including those affected by acute lymphoblastic leukemia.

The observed decrease in sensitivity in A549 cells may be attributed to a potentially higher level of endogenous asparagine biosynthesis in this cell type compared to K562 cells. This hypothesis is bolstered by previous investigations that have demonstrated that A549 cells exhibit substantial expression of ASNS, which can confer resistance to treatment with asparaginase [49]. ASNS inhibitors, such as bisabosqual A (Bis A), are capable of covalently modifying the K556 site on the ASNS protein. Bis A specifically targets ASNS, inhibiting the proliferation of A549 non-small-cell lung cancer cells in humans. When combined with L-asparaginase (L-ASNase), Bis A exhibits a synergistic effect [49]. Furthermore, there is a clear correlation between the level of ASNS activity and susceptibility to asparaginase-based therapy [8].

The literature data show that the different RrA mutants tested on the Jurkat cells largely lacked significant antitumor activity, except for RrA_{E149R, V150P}, which showed comparable activity to the commercial EcA Medac at dosages of 2 U/mL and above [38]. The native RrA also showed minimal activity against Jurkat cells. It is important to note that the cell lines used in the two datasets are different (K562 vs. Jurkat), which could influence the observed activity.

We propose several hypotheses regarding the enhanced cytostatic activity of RrA_{E149R, V150P, F151T}: (i) The close proximity of the mutations to the enzyme’s active site, potentially influencing catalytic turnover (V_max) and thus contributing to more efficient asparagine depletion. This is supported by the observed increase in catalytic efficiency. (ii) These mutations may contribute to the stabilization of the quaternary structure via enhanced electrostatic interactions. This structural stabilization is particularly relevant in the cellular environment, which has a high ionic strength that could otherwise destabilize the enzyme. (iii) The mutations in Mut3 enhance stability in the complex cellular milieu by conferring resistance to proteases to oxidation or other degradation factors present extracellularly or intracellularly (lysosomal enzymes) following internalization. Indeed, we observed a significantly higher resistance to trypsin exposure in Mut3 compared to the native RrA. This increased stability could result in a longer half-life and sustained asparagine depletion. Finally, the mutations in Mut3 could modulate interactions with cell surface receptors or transporters, potentially increasing internalization or improving delivery of the enzyme to its target site, thereby enhancing its cytostatic effect. This enhanced cellular uptake may also be linked to the observed stabilization of the tetrameric structure. We hypothesize that a near-neutral zeta potential and the presence of surface-exposed amino groups on the enzyme contribute to efficient cell penetration. This hypothesis aligns with previous findings demonstrating the clathrin-receptor-mediated internalization of enzymes [50].

In summary, the data highlight the differential sensitivity of various cancer cell lines to L-asparaginase treatment and indicate that RrA mutants can exhibit enhanced anti-leukemic activity compared to both the native enzyme and the commercial EcA preparation. The weaker effect observed in the A549 cells underscores the importance of considering cell-line-specific responses when evaluating the therapeutic potential of L-asparaginases.

3. Materials and Methods

3.1. Reagents

The following reagents were used in this work: L-asparagine monohydrate (BioChemica, Billingham, UK), trypsin (Sigma-Aldrich, St. Louis, MO, USA), sodium phosphate buffered saline solution (PBS; Eco-Service, St. Petersburg, Russia), salts, acids, alkalis, etc. (Reachim, Moscow, Russia).

3.2. Enzymes

(1) L-Asparaginase EcA from E. coli is a commercial drug manufactured by Medac (Germany) and Veropharm (Moscow, Russia).

(2) Native L-Asparaginase RrA from Rhodospirillum rubrum.

The RrA gene, in its native form, was isolated from the bacterium Rhodospirillum rubrum, which is part of the collection of the Department of Microbiology at Lomonosov Moscow State University in Moscow, Russia. This gene was obtained using the pET-23a vector from Novagen, Madison, WI, USA. To culture cells containing plasmids, 0.05 mg/mL of kanamycin was added to the medium. To induce the expression of the target protein, lactose was added to the expressed culture when the optical density at 600 nm (OD600) reached 1.9, with a final concentration of 0.2%. Cells were cultured for an additional 17–20 h and then centrifuged at 4000× g for 15 min. All steps of enzymatic purification were performed at a temperature of +4 °C. A sample of five grams of biomass was suspended in a buffer solution of 50 mL (20 mM PBS at pH 7.4 with the addition of 1 mM glycine and 1 mM EDTA). The mixture was then subjected to ultrasonic treatment. After that, cellular debris and intact cells were separated by centrifugation at 35,000× g for 30 min. The resulting supernatant, which contained the target enzyme, was applied to Q-sepharose columns manufactured by Pharmacia in Sweden. The protein was then eluted using a linear gradient ranging from 0 to 1.0 M NaCl. The fractions obtained were analyzed for protein content using absorption at 280 nm and enzyme activity measurements. Ultrafiltration, desalination, and replacement of the buffer were performed using Amicon membranes manufactured by Millipore in Burlington, Massachusetts. The samples were subsequently frozen and stored at a temperature of −20 degrees Celsius.

(3) Mutant forms of L-Asparaginase RrA from Rhodospirillum rubrum.

Mutant forms of RrA_{A64V, E67K} (RrA Mut1), RrA_{R118H, G120R} (RrA Mut2), and RrA_{E149R, V150P, F151T} (RrA Mut3) (Table 1) were obtained using site-directed mutagenesis (“Quik Change”) Single-Primer Reactions in Parallel using the pET23a vector (“Novagen”, Madison, WI, USA), carrying the RrA gene as a matrix.

The E. coli strains XL-blue and BL21(DE3), which had been transformed with the pET23 plasmid containing the RrA gene, were cultivated under the conditions previously described [32]. The synthesis of recombinant RrA was induced by adding lactose to the expression culture until the A600 density reached approximately 1.5, with a final concentration of 0.2%. After 14–20 h of induction, the cells were harvested by centrifugation at 3000 rpm for 5 min. The purification of recombinant L-asparaginase RrA and its mutants was achieved through the use of ultrasonic treatment and column chromatography with DEAE-Toyopearl 650M and QSepharose. The purified proteins were then desalted and concentrated on PM10 membranes (Millipore, Germany), stabilized with 0.5% glucose, and freeze-dried. The resulting enzymes were stored at −20 °C.

In Mut3, glutamic acid (E) was replaced by arginine (R) at position 149, valine (V) by proline (P) at position 150, and phenylalanine (F) by threonine (T) at position 151. In Mut1—alanine (A) replaces valine (V) in 64, and glutamic acid (E) replaces lysine (K) in 67. In Mut2—arginine (R) is replaced by histidine (H) at the 118th position and glycine (G) by arginine (R) at the 120th position. The N-fragment MASMTGGQQMGRGSSRQ is also present in the RrA Mut1 and Mut2. The mutations were confirmed by Sanger DNA sequencing.

The process of isolating and purifying the mutant forms of enzymes was carried out in the laboratory of medical biotechnology at the V. N. Orekhovich Institute of Biomedical Chemistry. The procedure involved the isolation and purification of L-asparaginases from the cells of a producing strain. This process included the use of ultrasound to treat the cell suspension, followed by the removal of debris through centrifugation. Subsequently, two stages of ion exchange chromatography were performed using columns filled with Q-Sepharose from Pharmacia (Uppsala, Sweden) and DEAE-Toyopearl 650 from ToyoSoda (Tokyo, Japan). Ultrafiltration, desalination, and concentration were also employed using PM10 membrane filters from Millipore (Billerica, MA, USA) in an Amicon cell. All stages of the purification process were conducted at +4 °C. The resulting L-asparaginase preparations were then stabilized with 0.5% glucose and lyophilized for storage at −80 °C.

The results of a 12% SDS-PAGE electrophoresis of the purified RrA variants are presented in the Supplementary Materials (Figure S1). The molecular mass of purified preparations corresponded to the theoretical values of 18 kDa (for truncated versions) or 19.8 kDa (for N17-fusions). The content of the target protein, calculated by Gel-Proanaluser 3.1.00.00 (Media Cybernetics, Rockville, MD, USA), represented 70−80% of the total protein (Figure S1).

3.3. FTIR Spectroscopy of Enzymes and Its Secondary Structure

The Fourier-transform infrared (FTIR) spectra of enzyme solutions were acquired using a MICRAN-3 FTIR microscope and a Bruker Tensor 27 spectrometer equipped with a liquid-nitrogen-cooled mercury cadmium telluride (MCT) detector. Deconvolution of the Amide I band (1600–1700 cm⁻¹) in the FTIR spectrum of asparaginases was performed to determine secondary structure content. This involved baseline correction, smoothing, and subsequent deconvolution using a curve-fitting algorithm (Gaussian functions) to resolve overlapping bands corresponding to different secondary structure elements (α-helices, β-sheets, turns, and random coils) taking into account the molar extinction coefficients of each component (α-helix 700 M⁻¹ × cm⁻¹, high-frequency β-sheet component (1695–1675 cm⁻¹) 180 M⁻¹ × cm⁻¹, major low-frequency β-sheet component (1639–1628 cm⁻¹) 180 M⁻¹ × cm⁻¹, and random coil structures 330 M⁻¹ × cm⁻¹) [51,52,53]. The relative areas under each resolved peak were then used to calculate the percentage contribution of each secondary structure element to the overall protein conformation.

3.4. Circular Dichroism (CD) Spectroscopy

Circular dichroism (CD) spectra of native L-asparaginase and its conjugates (with L-asparagine and L-aspartic acid) were recorded using a Jasco J-815 CD spectrometer (Jasco, Tokyo, Japan). Measurements were performed from 200–260 nm at 37 °C in a 300 µL quartz cuvette (1 mm path length). The spectra were obtained by averaging five scans with 1 nm increments. The samples were dissolved in 10 mM sodium phosphate buffer (pH 7.4) at protein concentrations ranging from 0.25 to 1.00 mg/mL.

3.5. Catalytic Activity Determination by CD Spectroscopy

L-asparaginase activity was determined using the Jasco J-815 CD spectrometer (Jasco, Tokyo, Japan). A solution of 40 mM L-asparagine in 10 mM sodium phosphate buffer (pH 7.4) was diluted to the desired concentration. An amount of 15 µL of a 1 mg/mL L-asparaginase solution (in the same buffer) was added to yield a final enzyme concentration of 0.05 mg/mL. The reaction was conducted at 37 °C in a 300 µL quartz cuvette (1 mm path length). The ellipticity at 210 nm was monitored over time. The hydrolysis rate was calculated using a calibration curve relating ellipticity to L-asparagine and L-aspartic acid concentrations, which were determined independently.

3.6. L-Asparaginase Thermodenaturation Parameter Determination

The thermodenaturation parameters were determined using the Jasco J-815 CD spectrometer. A solution of native enzyme or conjugate (0.5 mg/mL protein in 15 mM sodium phosphate buffer, pH 7.4) was heated from 20 °C to 100 °C at a rate of 1 °C/min in a 300 µL quartz cuvette (1 mm path length). Denaturation was monitored by following changes in ellipticity at 220 nm. Data were analyzed using JASCO Spectra Manager software (Jasco, Tokyo, Japan).

Considering the process:

(Native ↔ Denaturated), K = [N]/[D].

(1)

The obtained thermograms were approximated by an asymmetric sigmoid dependence in the Origin program (OriginLab Corporation, Northampton, MA, USA). Using the ratio: ΔG = −RT ln [(θn − θ)/(θ − θd)], where values of elliptic proteins in the native and denatured forms (θn and θd, respectively) were taken as the average values of enzyme ellipticity before and after the completion of the conformational transition of the protein.

Then, using the van’t Hoff isobar equation, ${\displaystyle \frac{dlnK}{dT}}={\displaystyle \frac{∆H}{R{T}^2}}$, the values of the effective enthalpy of denaturation were calculated.

The value of the effective entropy was found from the formula:

$$\Delta S={\displaystyle \frac{\Delta H-\Delta G}{T}}={\displaystyle \frac{\Delta H}{T}}+Rln[(\theta n-\theta )/(\theta -\theta d)]$$

3.7. Trypsinolysis Stability Assay

The stability of L-asparaginase and its modified forms during trypsinolysis was assessed using CD spectroscopy. Trypsin (0.01 mg/mL final concentration) was added to a solution of native or modified enzyme (0.5 mg/mL protein in pH 7.4 buffer). The mixture was incubated at 37 °C with samples taken at 5 min intervals (5, 10, 15, 20, 25, 30, 35, 40, 45, 50, 55, and 60 min). Residual catalytic activity was immediately determined by CD spectroscopy at 37 °C using reaction mixtures containing 0.05 mg/mL enzyme and 20 mM substrate in pH 7.5 buffer.

3.8. Cell Culture and Cytotoxicity Assay

K562 leukemia cells and A549 lung carcinoma cells were obtained from the Live Systems Collection (V. N. Orekhovich Institute of Biomedical Chemistry, Moscow, Russia). Cells were cultured in RPMI-1640 medium (Gibco, Thermo Fisher Scientific, Waltham, MA, USA) supplemented with 5% fetal bovine serum (Capricorn Scientific, Marburg, Germany) and 1% sodium pyruvate (Paneco) in a humidified incubator at 37 °C with 5% CO₂. Cytotoxicity was assessed using the MTT assay. The MTT assay protocol involved seeding cells at a defined density in 96-well plates. After 24 h, cells were treated with varying concentrations of the compounds under investigation. After 24 h, MTT solution (3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide) was added, and the cells were incubated for 12 h. The resulting formazan crystals were then dissolved using a solubilization DMSO solution, and the absorbance was measured using a microplate reader at wavelength 570 nm. Absorbance values were directly proportional to the number of viable cells.

3.9. FTIR Spectroscopy for Studying Enzyme-Cell Interactions

Cell suspensions (4–7 × 10⁵ cells/mL) were harvested by centrifugation (Eppendorf centrifuge 5415C, 2 × 5 min, 4000× g) and washed twice with sterile phosphate-buffered saline (PBS, pH 7.4). The cells were then resuspended in PBS to a concentration of 1 × 10⁷ cells/mL. A 20 µL aliquot of the cell suspension was deposited onto the spectrometer chamber, followed by the addition of 10 µL of enzyme solution (1–5 mg/mL). Samples were incubated at 37 °C, and the FTIR spectra were recorded at 5 min intervals over a one-hour period.

4. Conclusions

This study provides a comprehensive characterization of the conformational stability parameters, enzymatic activity, and anticancer effects of wild-type Rhodospirillum rubrum L-asparaginase (RrA) and three engineered mutants (Mut1—RrA_{A64V, E67K}; Mut2—RrA_{R118H, G120R}; Mut3—RrA_{E149R, V150P, F151T}), comparing them to commercially available EcA preparations. The mutations in this study aimed to reduce negative charge density near the subunit interface to stabilize the more active oligomeric form of the enzyme.

Our findings reveal significant improvements in several key parameters for the engineered RrA variants, suggesting promising avenues for developing improved L-asparaginase-based therapies. Structural changes in the RrA_{A64V, E67K} and RrA_{R118H, G120R} led to the improvement of kinetic parameters and enzyme stabilization due to the elimination of the steric hindrance in the case of this mutation upon oligomerization and increased protein stability. Mut3 focused on increasing antitumor activity. The N-terminal addition in Mut1 and Mut2 aimed for further biocatalytic improvements. These mutations were rationally selected based on previous research to potentially create synergistic effects leading to a superior enzyme.

The native RrA exhibited a V_max of 30 U/mg and a K_M of 4.5 ± 0.5 mM. Mut3 showed a remarkable increase in V_max to 57 U/mg, almost doubling its catalytic efficiency, while Mut2 demonstrated more moderate improvements. Importantly, all mutants showed increased K_M values (by 1–2 mM), suggesting some alterations in substrate binding affinity. However, the enhanced velocity in Mut3 outweighs this effect, leading to overall superior catalytic performance.

Thermal stability analysis revealed a critical difference between RrA and commercial EcA. Mut3 demonstrated markedly enhanced stability against trypsinolysis via a localized conformational change, retaining 45% of its activity after 30 min of exposure, significantly higher than native RrA (21%) and commercial EcAs (less than 10% remaining). Charge alteration, proline-induced rigidity, and hydrophobicity changes likely hinder trypsin access or reduce favorable binding.

Cytostatic activity against K562 leukemia cells confirmed the potential of the engineered RrA. Mut3 showed the most significant improvement, with an IC₅₀ of 10 U/mL, compared to native RrA (15 U/mL) and EcA-Veropharm (24 U/mL). Mut2 also demonstrated enhanced activity (IC₅₀ of 11.5 U/mL). These results suggest that the modifications in Mut2 (especially the G120R substitution) and Mut3 (V150P, in conjunction with E149R and F151T) led to a more effective interaction with cellular targets within K562 cells. A549 lung carcinoma cells showed consistently low sensitivity to all L-asparaginases, highlighting the specific effectiveness of this enzyme against leukemia cells. This low sensitivity may be attributed to a potentially higher level of endogenous asparagine biosynthesis in this cell type compared to K562 cells.

It was found that mutant substitutions of carboxyl groups to positively charged amino acid residues in the region of interunit contacts stabilize the oligomeric structure of the enzyme due to an improvement in the situation with a predominantly negative surface charge on RrA with limited positive charge clusters. This, in turn, plays a key role in the enzyme’s functioning and stability. The enhanced catalytic efficiency improved stability, and superior anticancer activity of Mut2, and especially Mut3, establish them as promising candidates for next-generation L-asparaginase-based therapies, opening avenues for more effective and less toxic treatments for leukemia.