Exploring Strategic Approaches for CvFAP Photodecarboxylation through Violet Light Irradiation

Abstract

In this study, we describe a light-driven photocatalytic decarboxylation of palmitic acid and related fatty acids using Chlorella variabilis fatty acid photodecarboxylase (CvFAP). By utilizing violet light emitting diode (LED) light (50 W; 397 nm), we achieved a remarkable conversion efficiency of 99% within just 4 min, surpassing the previous 79% conversion achieved in 60 min using blue LED light (300 W; 439 nm). Importantly, the use of 50 W violet LED light also resulted in a lower enzyme photoinactivation rate when compared to 300 W blue LED. Comparing the whole-cell biocatalyst with the enzymatic extract, we found that the former demonstrated superior catalytic performance and reduced susceptibility to photoinactivation. Furthermore, whole-cell biocatalyst reuse was demonstrated after five sequential batches. Employing this approach, we successfully synthesized 26 mmol L^-1 h^-1 of pentadecane, showcasing a promising strategy to improve productivity. These findings represent a significant advancement in CvFAP photodecarboxylation processes compared to the literature, utilizing an alternative light source, with potential implications to the biofuel sector.

Keywords:
photodecarboxylation; biocatalysis; green chemistry; violet light; CvFAP

Introduction

Sustainable alternatives, such as enzymes, for the chemical synthesis of molecules have been pursued to replace traditional chemical processes, which have long been associated with environmental harm.¹ In this context, photobiocatalysis has garnered significant attention over the past 25 years by combining the high selectivity and efficiency of biocatalysts with the transformative power of photoexcitation.²

The exploration of this captivating field led scientists to an important photoenzyme, Chlorella variabilis fatty acid photodecarboxylase (CvFAP), purified and identified for the first time in 2017.³ CvFAP possesses the capability to convert fatty acids into alka(e)nes in a light-driven reaction. The potential applications of this enzyme are remarkable,⁴ such as in the production of alkane-based biofuels and it has also been demonstrated in the synthesis of other compounds such as chiral secondary fatty alcohols.⁵ Mutant variants of CvFA P have also been proposed and studied,⁶,⁷ such as other enzymes of FAP group⁸,⁹ and possible ancestral decarboxylases.¹⁰

The literature surrounding CvFAP has already reported the dependence of light intensity and wavelength in the reaction yields.³,¹¹,¹²,¹³ Benincá et al.¹¹ described the improvement in the conversion of palmitic acid to pentadecane by increasing the power of the blue light emitting diode (LED). Furthermore, the use of high-power LEDs (blue and white) was demonstrated in continuous flow systems reducing the reaction time and the amount of enzymatic extract and achieving 99% conversion in 10 min. The benefit of continuous flow technology was then proved for CvFAP mediated photodecarboxylation reactions.¹¹

Although blue LEDs (ca. 430-490 nm) are the most commonly used light source, the reported high photoinactivation by the blue light is a concern.¹²,¹⁴,¹⁵ Green light (ca. 495-570 nm) has been reported to enhance the photostability of CvFAP, offering a potential advantage in certain applications.¹⁶ However, it is noteworthy that this enhancement in photostability is accompanied by a tradeoff in terms of overall performance.¹²,¹⁶

França et al.¹⁷ recently showed that the sunlight is a great alternative to the blue LEDs in terms of efficiency. Full conversion (> 99%) of palmitic acid to pentadecane was attained in just 12 min compared to > 99% conversion obtained after 50 min with blue LED,¹¹ despite the lower light intensity presented by the sunlight in the visible range (380-700 nm). Therefore, the present work followed our previous investigation of the CvFAP photodecarboxylation, and herein, we present our results on the use of violet LED (397 nm) as an alternative light source for decarboxylation reactions.

Experimental

Solvents and reagents

Tris base (Sigma-Aldrich, Burlington, Massachusetts, USA), HCl 37% (Vetec Química, Duque de Caxias, Brazil), Kanamycin (Fluka, Buchs, Switzerland), methyl sulfoxide (Tedia, USA), ethyl acetate (Tedia, USA), ethanol (Vetec Química), methanol (Vetec Química, Duque de Caxias, Brazil), sodium sulfate anhydrous (Vetec Química, Duque de Caxias, Brazil), palmitic acid (Sigma-Aldrich, Burlington, Massachusetts, USA), Pierce™ BCA Protein Assay Kit (ThermoFisher Scientific, Waltham, Massachusetts, USA) were used in the present work.

Equipment

Analytical balance Mettler Toledo (USA), magnetic stirrer heating plate Model 753A-FISATOM (Brazil), orbital shaker incubator Marconi (Brazil), gas chromatographyflame ionization detection (GC-FID) Hewlett-Packard (USA) Model GC-6890C, Shimadzu (Japan) GC2014 GC-FID-Cpsil 5 CB column (50 m × 0.53 mm × 1.0 µm), portable digital HighMed KR812 luximeter until 200000 lux (Brazil), ultrasonic sonicator VIBRA-CELL VCX 500 Sonics USA) were used in the present work.

Strain, vector and materials

Escherichia coli DH5α strains were employed for deoxyribonucleic acid (DNA) manipulation purposes. The plasmid housed the CvFAP gene as described in previous work.¹¹ Recombinant enzyme production involved transforming competent E. coli BL21 (DE3) cells with this plasmid. Kanamycin, obtained from Fluka (Buchs, Switzerland), was used in the process. For experimental requirements, palmitic acid was purchased from Sigma-Aldrich (Burlington, Massachusetts, USA). Additionally, the Pierce™ BCA Protein Assay Kit was purchased from ThermoFisher Scientific (Waltham, Massachusetts, USA).

Preparation of the biocatalysts

The methodology for enzyme expression followed the protocol outlined by Huijbers et al.¹⁸ Following CvFAP expression, E. coli BL21 (DE3) cells were stored and used to prepare the enzymatic extract (CvFAP_EE) following previous protocols of the research group.¹¹ The total protein concentration of the CvFAP_EE samples was determined and standardized using the Pierce™ BCA Protein Assay Kit, following the supplier’s instructions. The whole-cell biocatalyst (CvFAP_WC) was prepared by resuspension in reaction buffer (100 mM Tris-HCl buffer, pH 8.5, containing 1 mM phenylmethylsulfonyl fluoride (PMSF) and 5% glycerol) with 11 mg mL^-1 of dry cell weight, measured using spectrophotometric analysis at a wavelength of 600 nm.

General photocatalytic reactions in batch with LED light

Standard photoenzymatic decarboxylation reactions were conducted in triplicate as previously reported by Benincá et al.¹¹ containing CvFAP_EE or CvFAP_WC as biocatalysts. Different LED lights were used in the reactions, such as a 300 W blue LED lamp and a 50 W violet LED lamp. The reusability tests were performed for CvFAP_WC biocatalysts by starting a standard batch reaction using a 50 W violet LED lamp for 15 min. Then, the cells were centrifuged at 6.000 rpm for 15 min at room temperature and recovered to subsequent batches. Photostability tests were performed by pre-illuminating the CvFAP_WC and CvFAP_EE biocatalysts in the absence of the substrate using different illumination times and light sources. Then, the substrate was added, and 1 h standard batch reactions were performed using the same light source used for the pre-illumination.

Gas chromatography analysis

Samples were prepared in ethyl acetate with a solvent-to-reaction ratio of 3:1 and subsequently dried using Na₂SO₄. Chromatography areas were used to determine conversion percentages using the Shimadzu GC2014 GC-FID-Cpsil 5 CB column (50 m × 0.53 mm × 1.0 µm) as reported by Benincá et al.¹¹

Results and Discussion

Our previous research¹⁷ findings have highlighted the promising potential of sunlight for CvFAP photodecarboxylation. In this way, the approach using the sunlight has motivated us to a more specific investigation aiming to explore the correlation between the efficacy of our results with the broad emission spectra of sunlight. The UV-Vis absorption spectrum of the purified CvFAP-bound flavin adenine dinucleotide (FAD) cofactor and free FAD exhibits maxima at 467 nm (blue) and also at 389 nm (violet).¹⁵ Typically, the blue range is used for photoactivation purposes. However, a previous study¹³ has reported the utilization of low-power strip LEDs (1-1.4 W) emitting violet light and showed similar results to blue light under these experimental conditions.

We propose a batch study utilizing a 50 W violet LED lamp (397 nm) with a reaction time of up to 40 min (Tables S1 and S2, ^{Supplementary Information (SI)} section). Remarkably, complete conversion (> 99%) was achieved within mere 4 min. Consequently, we compared the results using 300 W blue LED lamps with the performance of the violet LED (397 nm) under the same experimental conditions (Table S3, SI section). References from the literature were also included to highlight the differences between light sources. Clearly, the 50 W violet LED exhibits superior efficiency compared to the 300 W blue LED, requiring significantly less power output and light intensity (Table S6, SI section) to achieve a similar conversion for the same substrate.

The discrepancy observed in the data obtained by Winkler et al.¹³ (44% conversion for blue light and 41% for violet light in 20 min) might be attributed to variations in enzyme concentration and specific form, the utilization of lower-power LEDs with slightly different wavelengths and specific photoreactor. Similarly notable discrepancies were observed between the results obtained from experiments conducted with low-power strip LEDs emitting blue light¹³ and our previous research findings involving high-power blue LEDs.¹¹ By increasing the intensity of blue light, we were able to enhance the reaction rate under our experimental conditions,¹¹ aligning with the observations reported by Sorigué et al.³ using white light. However, according to Winkler et al.,¹³ a plateau effect was observed after reaching 20% of the maximum intensity tested for blue light.

A broad range of long-chain fatty acids was converted by CvFAP using high-power blue LED light source, in up to 1 h reaction time (Table S4, SI section). It is clear the preference of CvFAP to saturated fatty acid chains higher than 14 carbons. Unsaturated fatty acids could also lead to reasonable results while C14 fatty acids allowed only 23% of conversion after 1 h. Those results were comparable to 14 h reactions using higher substrate concentration (30 mM) reported by Huijbers et al.,¹⁸ with slightly higher conversion obtained for linoleic acid.

Herein, we specifically focused on exploiting the potential of CvFAP in the form of enzymatic extract (CvFAP_EE). However, CvFAP has been reported in several other forms such as cell-free extract (which involves centrifugation following cell lysis), purified enzyme and whole cell.¹⁸,¹⁹ Considering the advantages associated with whole cells, including the elimination of cell disruption and simplification of the experimental process, we performed time-course reaction analysis using CvFAP in the form of whole cells (CvFAP_WC). Notably, the achieved conversions at a concentration of only 11 mg mL^-1 of dry cell weight (Table 1) were comparable to the results obtained using enzymatic extract with a total protein concentration of 2.8 mg mL^-1 (Table S2, SI section).

Table 1
Palmitic acid photodecarboxylation reaction mediated by CvFAP_WC using a 50 W violet LED lamp (397 nm) with different initial substrate concentrations for 60 min

time / min	Conversiona / %
	13 mM	26 mM	65 mM
5	95 ± 0.2	-	-
10	> 99 ± 0.2	93 ± 0.2	12 ± 5
60	-	> 99 ± 0.2	47 ± 0.4

• ^a
  The conversions were measured by gas chromatography and calculated by integration of chromatogram peak area. Reaction conditions: to a transparent flask reactor (total volume 1 mL) different concentrations of palmitic acid (13, 26 and 65 mM) and Tris-HCl buffer (pH 8.5, 100 mM) 30% dimethyl sulfoxide (DMSO), containing the CvFAP_WC (11 mg mL^-1 of dry cell weight) were added. The reaction was carried out at 37 °C for 10 min using a 50 W violet LED (397 nm).

Guo et al.¹⁹ conducted a thorough comparison between CvFAP whole-cells and enzymatic extract biocatalysts, resulting in similar global conversion rates, with 15.2 mmol L^-1 h^-1 achieved by the whole-cell biocatalyst at a concentration of 35.2 mg mL^-1 of dry cell weight. It is noteworthy that higher conversions were achieved at the present work with significantly less biocatalyst (11 mg mL^-1 of dry cell weight) in reduced time, as indicated in Table 1. Moreover, a significant decline in conversion rates was observed in our previous photodecarboxylation studies using higher substrate concentrations illuminated by blue LED¹¹ (16% conversion at 39 mM) and sunlight¹⁷ (36% conversion at 34.5 mM) with the enzymatic extract for 1 h. However, it is worth mentioning that even with the increased substrate concentration of 65 mM, we were able to achieve an excellent conversion rate of 30 mmol L^-1 h^-1 palmitic acid within just 1 h using violet LED and whole-cell biocatalysts.

The reusability of whole cell biocatalysts is a highly advantageous feature, offering potential cost reductions and improved process efficiency.²⁰ The extent of reusability and the optimal conditions for biocatalyst reuse depend on several factors, including the specific biotransformation reaction and biocatalyst stability. In our study, we assessed the reusability of the CvFAP_WC biocatalyst under 50 W violet LED illumination and observed exceptional performance, with a consistent conversion rate of 75% maintained even after five consecutive batch cycles, although the use of longer reaction times was required in most cases (Table 2).

Table 2
Conversions obtained for whole-cell biocatalyst reuse in photodecarboxylation reactions using a 50 W violet LED light and centrifugation to recover the cells

Batch	Reaction time / min	Conversiona / %
1	15	> 99 ± 0.2
2	30	> 99 ± 0.2
3	40	> 99 ± 0.2
4	40	97 ± 0.2
5	40	75 ± 0.4

• ^a
  The conversions were measured by gas chromatography and calculated by integration of chromatogram peak area. Reaction conditions: to a transparent flask reactor (total volume 1 mL), palmitic acid (13 mM) and Tris-HCl buffer (pH 8.5, 100 mM) 30% DMSO, containing the CvFAP_WC (11 mg mL^-1 of dry cell weight), were added. The reaction was conducted at 37 °C using a 50 W violet LED (397 nm) for 15 min in batch 1. The cells were then centrifuged (6000 rpm; 15 min; room temperature) to be reused under the same reaction conditions for each subsequent batch with 30-min reaction in batch 2, and 40-min reactions in batches 3, 4, and 5.

Photostability is a relevant concern regarding CvFAP photodecarboxylation and has been investigated by various research groups.¹⁴,¹⁵,¹⁸ Wu et al.¹⁴ have shown that excessive light exposure decreases CvFAP activity, especially in the absence of free fatty acids (FFA). The photostability of CvFAP is influenced by light intensity and oxygen presence,¹²,¹⁴,²¹ such as the biocatalyst type, with cell-free extracts remarkably more stable than the purified enzyme.¹⁴,¹⁸ Additionally, the spectral window of irradiation plays a significant role, as exemplified for CvFA P ¹⁶ and the photodecarboxylase from Micractinium conductrix (McFAP).⁸ High stability was demonstrated by McFAP in the dark and under red light pre-illumination, 40% residual activity was observed after 3 h of sunlight pre-illumination, while complete inactivation occurred after 30 min under blue light. Green light has been used for photodecarboxylation reactions using CvFAP with lower catalytic activity and higher photostability.¹²,¹⁶

Thus, the photostability of CvFAP biocatalysts (CvFAP_EE and CvFAP_WC) was evaluated by the pre-illumination of the biocatalysts under the two light sources, 300 W blue LED and 50 W violet LED. The results demonstrated higher stability when using whole-cells biocatalysts compared to enzymatic extract for both light sources (Table 3). Previous reports¹⁸ have also indicated that the addition of cell-free extract to the purified enzyme enhances stability. The encapsulation of the enzyme of interest within the whole-cells provides a more favorable environment,²² potentially facilitating the presence of fatty acids to maintain the active site fulflled.¹⁴

Table 3
Conversions obtained for 1 h CvFAP photodecarboxylation reactions after the pre-illumination of the biocatalysts using different light sources

Light source	Biocatalyst	Pre-illumination time
		15 min	30 min	40 min
		Conversion / %
Blue LED300 W	enzymatic extract	45 ± 0.4	16 ± 5.8	0
	whole-cell	71 ± 0.3	20 ± 0.4	11 ± 5.4
Violet LED50 W	enzymatic extract	77 ± 0.3	35 ± 0.4	8 ± 5
	whole-cell	90 ± 0.2	71 ± 0.3	55 ± 1.8

• Reaction conditions: to a transparent flask reactor, the CvFAP_EE (7.1 mg mL^-1 of total protein) or CvFAP_WC (11 mg mL^-1 of dry cell weight) were added in Tris-HCl buffer (pH 8.5, 100 mM) and incubated for 15, 30 and 40 min. Palmitic acid (13 mM) dissolved in DMSO (30%) was then added and the reactions were carried out at 37 ºC for 1 h using a 50 W violet LED (397 nm) or a 300 W blue LED (439 nm). The conversions were measured by gas chromatography.

Both CvFAP biocatalysts exhibited higher stability under 50 W violet LED light, which can be attributed to the lower energy power and the differences in light intensity and wavelength compared to the 300 W blue LED light. The possible accumulation of superoxide radical and singlet oxygen under high power blue LED illumination²¹ may explain the results for these light sources.

The remarkable efficiency of violet LEDs compared to blue LEDs demonstrated in the present work opens a new window of opportunities for CvFAP mediated reactions. The use of violet LED can not only increase the reaction yield and photostability of the biocatalyst but also represents a path to unveil new aspects of its catalytic mechanism. Further analysis using spectrophotometric methods and reactions in the absence of oxygen could provide a rationale for our experimental observations, such as previously demonstrated for blue and green light illumination.¹⁶,²¹

Conclusions

In conclusion, these studies reinforce the importance of wavelength on decarboxylation mediated by CvFA P and corroborate the previous reports using the broad electromagnetic spectrum of sunlight. The present work has shown that CvFAP photodecarboxylation reaction is possible to be performed under high power violet LED light with very fast reaction conditions where full conversion can be obtained after only 4 min. To the best of our knowledge, the present report also shows the best conversion rates and pentadecane concentrations obtained for CvFAP photodecarboxylation of palmitic acid using the strategic whole-cells biocatalysts.

Acknowledgments

The authors acknowledge the National Council for Scientific and Technological Development (Conselho Nacional de Desenvolvimento Científico e Tecnológico-CNPq) and Carlos Chagas Filho Foundation for Support of Research of the State of Rio de Janeiro (Fundação Carlos Chagas Filho de Amparo à Pesquisa do Estado do Rio de Janeiro-FAPERJ) for financial support. We also thank FAPESP-São Paulo State Research Foundation (K.T.O. grants 2013/07276-1, 2019/27176-8, 2020/06874-6 and 2023/0402-8). This study was financed in part by the Coordenação de Aperfeiçoamento de Pessoal de Nível Superior-Brasil (CAPES)-Finance Code 001.

Supplementary Information

Supplementary information is available free of charge at http://jbcs.sbq.org.br as PDF file.

References

• ¹
  Sheldon, R. A.; Woodley, J. M.; Chem. Rev. 2018, 118, 801. [Crossref]
  » Crossref
• ²
  Emmanuel, M. A.; Bender, S. G.; Bilodeau, C.; Carceller, J. M.; DeHovitz, J. S.; Fu, H.; Liu, Y.; Nicholls, B. T.; Ouyang, Y.; Page, C. G.; Qiao, T.; Raps, F. C.; Sorigué, D. R.; Sun, S. Z.; Turek-Herman, J.; Ye, Y.; Rivas-Souchet, A.; Cao, J.; Hyster, T. K.; Chem. Rev. 2023, 123, 5459. [Crossref]
  » Crossref
• ³
  Sorigué, D.; Légeret, B.; Cuiné, S.; Blangy, S.; Moulin, S.; Billon, E.; Richaud, P.; Brugière, S.; Couté, Y. ; Nurizzo, D.; Müller, P.; Brettel, K.; Pignol, D.; Arnoux, P.; Li-beisson, Y.; Peltier, G.; Beisson, F.; Plant Sci. 2017, 907, 903. [Crossref]
  » Crossref
• ⁴
  Guo, X.; Xia, A.; Zhang, W.; Huang, Y.; Zhu, X.; Zhu, X.; Liao, Q.; Bioresour. Technol. 2023, 367, 128232. [Crossref]
  » Crossref
• ⁵
  Zhang, W.; Lee, J. H.; Younes, S. H. H.; Tonin, F.; Hagedoorn, P. L.; Pichler, H.; Baeg, Y. ; Park, J. B.; Kourist, R.; Hollmann, F.; Nat. Commun. 2020, 11, 2258. [Crossref]
  » Crossref
• ⁶
  Santner, P.; Szabó, L. K.; Nahuel, C. S.; Merrild, A. H.; Hollmann, F.; Kara, S.; Eser, B. E.; ChemCatChem 2021, 13, 4038. [Crossref]
  » Crossref
• ⁷
  Li, D.; Han, T.; Xue, J.; Xu, W.; Xu, J.; Wu, Q.; Angew. Chem., Int. Ed. 2021, 60, 20863. [Crossref]
  » Crossref
• ⁸
  Ma, Y. ; Zhong, X.; Wu, B.; Lan, D.; Zhang, H.; Hollmann, F.; Wang, Y.; Chin. J. Catal. 2023, 44, 160. [Crossref]
  » Crossref
• ⁹
  Zeng, Y.; Yin, X.; Liu, L.; Zhang, W.; Chen, B.; Mol. Catal. 2022, 532, 112717. [Crossref]
  » Crossref
• ¹⁰
  Sun, Y. ; Calderini, E.; Kourist, R.; ChemBioChem 2021, 22, 1833. [Crossref]
  » Crossref
• ¹¹
  Benincá, L. A. D.; França, A. S.; Breda, G. C.; Le, R. A. C.; Almeida, R. V.; Hollmann, F.; de Souza, R. O. M. A.; Mol. Catal. 2022, 528, 112469. [Crossref]
  » Crossref
• ¹²
  Simić, S.; Jakštaitė, M.; Huck, W. T. S.; Winkler, C. K.; Kroutil, W.; ACS Catal. 2022, 12, 14040. [Crossref]
  » Crossref
• ¹³
  Winkler, C. K.; Simic, S.; Jurkas, V. ; Bierbaumer, S.; Schmermund, L.; Poschenrieder, S.; Berger, S. A.; Kulterer, E.; Kourist, R.; Kroutil, W.; ChemPhotoChem 2021, 5, 957. [Crossref]
  » Crossref
• ¹⁴
  Wu, Y.; Paul, C. E.; Hollmann, F.; ChemBioChem 2021, 22, 2420. [Crossref]
  » Crossref
• ¹⁵
  Lakavath, B.; Hedison, T. M.; Heyes, D. J.; Shanmugam, M.; Sakuma, M.; Hoeven, R.; Tilakaratna, V.; Scrutton, N. S.; Anal. Biochem. 2020, 600, 113749. [Crossref]
  » Crossref
• ¹⁶
  Xia, A.; Guo, X.; Chai, Y.; Zhang, W.; Huang, Y. ; Zhu, X.; Zhu, X.; Liao, Q.; Chem. Commun. 2023, 59, 6674. [Crossref]
  » Crossref
• ¹⁷
  França, A. S.; Breda, G. C.; de Oliveira, K. T.; Almeida, R. V.; Hollmann, F.; de Souza, R. O. M. A.; Front. Catal. 2023, 3, 1165079. [Crossref]
  » Crossref
• ¹⁸
  Huijbers, M. M. E.; Zhang, W.; Tonin, F.; Hollmann, F.; Angew. Chem., Int. Ed. 2018, 57, 13648. [Crossref]
  » Crossref
• ¹⁹
  Guo, X.; Xia, A.; Li, F.; Huang, Y. ; Xianqing, Z.; Zhang, W.; Zhu, X.; Liao, Q.; Energy Convers. Manage. 2022, 255, 115311. [Crossref]
  » Crossref
• ²⁰
  de Carvalho, C. C. C. R.; Microb. Biotechnol. 2017, 10, 250. [Crossref]
  » Crossref
• ²¹
  Guo, X.; Xia, A.; Zhang, W.; Li, F.; Huang, Y.; Zhu, X.; Zhu, X.; Liao, Q.; Chin. Chem. Lett. 2022, 34, 107875. [Crossref]
  » Crossref
• ²²
  Madavi, T. B.; Chauhan, S.; Keshri, A.; Alavilli, H.; Choi, K. Y. ; Pamidimarri, S. D. V. N.; Biofuels, Bioprod. Biorefin. 2022, 16, 859. [Crossref]
  » Crossref

Edited by

Editor handled this article: Brenno A. D. Neto

Publication Dates

• Publication in this collection
  22 Mar 2024
• Date of issue
  2024

History

• Received
  11 Dec 2023
• Accepted
  01 Mar 2024