species to be quite easy [167]. Thus, spontaneous formation of this halogen bond in C-3 from an epoxide intermediate could also be an option for these compounds.

<figure><img src="image_31.png" /><figcaption>**Figure 28.** Metabolites present in cluster 4, aranochlor A (63) from *Pseudoarachniotus* sp., dankastatin A (64) and B (65) from *Gymnascella* sp.</figcaption></figure>

Interesting metabolites from basidiomycetes can be found in cluster 5 like pterulone (66) produced by *Pterula* sp. (Figure 29) [168]. These metabolites carry a chlorine atom on a terminal double bond which is unusual. This cluster gathers 9 metabolites like calocerin B (67) or 2,3-dihydro-1-benzoxepin derivative 4a (68) both produced by *Favolaschia* sp. [169]. The halogenation takes place on a *sp*² carbon, but no enzyme described before seems to be able to perform this type of reaction. Indeed, FDHs and haloperoxidases catalyze electrophilic halogenation. Here, the total synthesis of pterulone (66) has been described using the way of nucleophilic addition. A Wittig reaction was used to introduce chlorine on a ketone using (chloromethyl)triphenylphosphonium chloride with n-butyllithium in THF at 0 °C, which corresponds to harsh chemical conditions [170]. So far, no enzyme performing this kind of reaction in fungi has been described to selectively halogenate ketones by displacing an oxygen atom. Further studies on the biosynthetic pathways of these compounds would be needed to further understand the way they are naturally halogenated.

<figure><img src="image_32.png" /><figcaption>**Figure 29.** Some metabolites from cluster 5: pterulone (66), calocerin (67), and 2,3-dihydro-1-benzoxepin derivative 4a (68).</figcaption></figure>

### 3.4. Compounds in the Haloalkane Class

With the exception of caldariomycin (2) and the very particular diaporthin derivatives (43, 44 and 45) mentioned above, all clusters for which a halogenating enzyme was described corresponded to haloarenes or haloalkene molecules with a FDH involved. From this observation, molecules classified as haloalkanes (or alkyl halides) occurred to be of interest considering their halogenation mechanism. In fact, preparation of alkyl halides in organic chemistry usually requires the use of radical reactions on alkanes or alkenes, with reagents such as *N*-bromosuccinimide (NBS), *N*-chlorosuccinimide (NCS), Br₂ or Cl₂ and energy such as light or heat. The formation of the carbon-halogen bonds on *sp*³ carbons can be difficult and require harsh chemistry. If these types of bonds are present in nature, they might be catalyzed by enzymes. In relation with what was previously presented, non-heme
iron-dependent halogenases (NHFe) and S-adenosylmethionine-dependent halogenases (SAM-H) are the two types of halogenation enzymes that are capable of carrying out halogenations on non-activated aliphatic chains, using radical or nucleophilic mechanisms. However, these enzymes are poorly known and described in fungi. This is why a closer look into the structures in the haloalkane class was carried out.

Among molecules classified as haloalkanes, the previously mentioned diaporthin derivatives could be retrieved in cluster 6, which gathered 9 molecules produced by different species of fungi such as *Hamigera* sp., *Ampelomyces* sp., *Penicillium* sp. and another undescribed species (Figure 30). They correspond to different isocoumarins classified as haloalkanes. The halogen atoms are found on the aliphatic chains carried by the molecules. AoiQ, which is a particular multitask enzyme with both FDH and SAM motifs, is involved in the biosynthesis of (43), (44) and (45) as explained above. Dichlorodiaportin (44) is dichlorinated by this enzyme but also methylated to obtain the methoxy groups. Regarding the similarity of structure here, it can be assumed that a similar protein is involved in the biosynthesis of (9R)-8-methyl-9,11-dichlorodiaportin (70) [171], peniisocoumarin F and J (71, 72) [172] and all the metabolites present in this cluster with halogenated unactivated aliphatic chains.

<figure><img src="image_33.png" /><figcaption>**Figure 30.** Metabolites displaying chlorination on aliphatic chains in the studied cluster 6, peniisocoumarin C (69) from *Penicillium commune*, (9R)-8-methyl-9,11-dichlorodiaportin (70) from *Hamigera fusca*, and peniisocoumarin F (71) and J (72) from *Penicillium commune*.</figcaption></figure>

If no enzyme could be clearly proposed in this kind of structure, halogenation on aliphatic chains seem to be complicated with weak reactivity. However, all molecules present a hydroxyl group on the carbon in α position. This suggests the possible spontaneous formation of halogen bonds here from an epoxide intermediate again. In fact, peniisocoumarin C (69), which was isolated together with peniisocoumarins A-B and D-J from *Penicillium commune* actually corresponds to the proposed intermediate allowing good reactivity for spontaneous halogenation on that position [173,174].

Further considering haloalkane-type molecules, cluster 7 attracted our attention as it included halogenated secondary metabolites belonging to the class of non-ribosomal peptides, described from strains of the genera *Penicillium* sp., *Talaromyces* sp., *Beauveria* sp. and an unknown strain (Figure 31). Once again, halogens are placed on saturated carbon chains and therefore on unreactive positions. In this case, we can refer to the cyclochlorotine (73) biosynthesis reported in 2016 [175]. Considering the chlorine atoms on the molecules, the authors suggested the incorporation of a 3,4-dichloroproline during the formation of the peptide. However, they could not find any sequence coding for a halogenase in the BGC of cyclochlorotine (73). Despite their attempts to identify elsewhere in the genome the halogenase responsible for the formation of the unusual Pro(Cl₂), by searching for conserved signatures of NHFe halogenases and α-ketoglutarate dependent halogenases, they could not detect any. They also hypothesized the combined action of a dehydrogenase and a FDH, but overexpression experiments of candidate genes were not
iron-dependent halogenases (NHFe) and S-adenosylmethionine-dependent halogenases (SAM-H) are the two types of halogenation enzymes that are capable of carrying out halogenations on non-activated aliphatic chains, using radical or nucleophilic mechanisms. However, these enzymes are poorly known and described in fungi. This is why a closer look into the structures in the haloalkane class was carried out.

Among molecules classified as haloalkanes, the previously mentioned diaporthin derivatives could be retrieved in cluster 6, which gathered 9 molecules produced by different species of fungi such as *Hamigera* sp., *Ampelomyces* sp., *Penicillium* sp. and another undescribed species (Figure 30). They correspond to different isocoumarins classified as haloalkanes. The halogen atoms are found on the aliphatic chains carried by the molecules. AoiQ, which is a particular multitask enzyme with both FDH and SAM motifs, is involved in the biosynthesis of (43), (44) and (45) as explained above. Dichlorodiaportin (44) is dichlorinated by this enzyme but also methylated to obtain the methoxy groups. Regarding the similarity of structure here, it can be assumed that a similar protein is involved in the biosynthesis of (9R)-8-methyl-9,11-dichlorodiaportin (70) [171], peniisocoumarin F and J (71, 72) [172] and all the metabolites present in this cluster with halogenated unactivated aliphatic chains.

<figure><img src="image_33.png" /><figcaption>**Figure 30.** Metabolites displaying chlorination on aliphatic chains in the studied cluster 6, peniisocoumarin C (69) from *Penicillium commune*, (9R)-8-methyl-9,11-dichlorodiaportin (70) from *Hamigera fusca*, and peniisocoumarin F (71) and J (72) from *Penicillium commune*.</figcaption></figure>

If no enzyme could be clearly proposed in this kind of structure, halogenation on aliphatic chains seem to be complicated with weak reactivity. However, all molecules present a hydroxyl group on the carbon in α position. This suggests the possible spontaneous formation of halogen bonds here from an epoxide intermediate again. In fact, peniisocoumarin C (69), which was isolated together with peniisocoumarins A-B and D-J from *Penicillium commune* actually corresponds to the proposed intermediate allowing good reactivity for spontaneous halogenation on that position [173,174].

Further considering haloalkane-type molecules, cluster 7 attracted our attention as it included halogenated secondary metabolites belonging to the class of non-ribosomal peptides, described from strains of the genera *Penicillium* sp., *Talaromyces* sp., *Beauveria* sp. and an unknown strain (Figure 31). Once again, halogens are placed on saturated carbon chains and therefore on unreactive positions. In this case, we can refer to the cyclochlorotine (73) biosynthesis reported in 2016 [175]. Considering the chlorine atoms on the molecules, the authors suggested the incorporation of a 3,4-dichloroproline during the formation of the peptide. However, they could not find any sequence coding for a halogenase in the BGC of cyclochlorotine (73). Despite their attempts to identify elsewhere in the genome the halogenase responsible for the formation of the unusual Pro(Cl₂), by searching for conserved signatures of NHFe halogenases and α-ketoglutarate dependent halogenases, they could not detect any. They also hypothesized the combined action of a dehydrogenase and a FDH, but overexpression experiments of candidate genes were not
conclusive. Finally, one recent study on cyclochlorotine (73) BGC in *P. islandicum* found that 73 is dichlorinated by a tailoring process involving an enzyme named CctP2, which can halogenate unactivated carbons. This enzyme was described to share no homology with any known halogenases, leading it to be a new type of enzyme that activates inert C-H bonds like NHFe halogenases [176].

<figure><img src="image_34.png" /><figcaption>**Figure 31.** Example of halogenated non ribosomal peptides found in this specific cluster 7, cyclochlorotine (73) and cyclochlorotine B (74) from *Penicillium islandicum*, destruxin-A4-cyclohydrin (75) from unknown fungi.</figcaption></figure>

For monochlorinated compounds in this cluster, while spontaneous hydrohalogenation involving hydrohalic acid (HCl, HBr) [177] on a double bond can still be considered as an explanation for their formation, we can assume similar halogenation mechanisms inside a family of compounds.

To finish, as already shown, halogenation of alkanes is well represented in fungi. NHFe halogenases and SAM-dependant halide methyltransferases (like AoiQ) are the only known enzymes that can halogenate unactivated aliphatic carbons. However, they are poorly described in fungi. And yet, many fungal compounds carry halogen on aliphatic chains as evidenced in clusters 8, 9 and 10.

Cluster 8 is composed of curtachalasin L (76), I (77) and K (78) together with xylarichalasin A (79) produced by *Xylaria* sp. [178,179] (Figure 32). All these metabolites carry halogenations on cyclohexane rings. Interestingly, during 12-epi-fischerindole G formation catalyzed by the bacterial NHFe halogenases WelO5 in *Hapalosiphon welwischii*, halogenation of a cyclohexane has been described [138]. We can then hypothesize that a similar protein is produced by this fungus and involved in the biosynthesis of these compounds. This hypothesis can be applied for metabolites in cluster 9 where halogenation again occurs on the cyclohexane ring of the three metabolites pestalotiopens A (80), B (81) and C (82) produced by *Pestalotiopsis* sp. [180] (Figure 32). Our search for WelO5 homologous proteins in Xylariales genomes available on MycoCosm gave no result. However, it is possible that even if a fungal NHFe halogenase was present, its sequence might be too different from bacterial NHFe halogenase WelO5.

Cluster 10 is composed of the ribosomally synthesized and post-translationally modified peptides (RiPPs) victorins B (83), D (84) and E (85) from *Cochliobolus victoriae* [181–183] (Figure 33). All these molecules present terminal chlorines on aliphatic chains. The chlorinated lipopeptide barbamide described in the literature from the marine cyanobacterium *Lyngbya majuscula* present the same trichlorinated carbon sp³ as 84. Biosynthesis of barbamide has been investigated and was shown to be performed by a hybrid NRPS/PKS enzyme. In the corresponding biosynthetic cluster, barB1 and barB2 encode two proteins that share homology with NHFe halogenases SyrB1-SyrB2 and CmA B from *Pseudomonas syringae*. These enzymes have been described to halogenate the amino-acid L-threonine for SyrB2 and L-alloisoleucine for CmA B before their insertion into non ribosomal peptides (NRP) [139,184]. For barbamide, trichlorination of leucine was then shown to be catalyzed
conclusive. Finally, one recent study on cyclochlorotine (73) BGC in *P. islandicum* found that 73 is dichlorinated by a tailoring process involving an enzyme named CctP2, which can halogenate unactivated carbons. This enzyme was described to share no homology with any known halogenases, leading it to be a new type of enzyme that activates inert C-H bonds like NHFe halogenases [176].

<figure><img src="image_34.png" /><figcaption>**Figure 31.** Example of halogenated non ribosomal peptides found in this specific cluster 7, cyclochlorotine (73) and cyclochlorotine B (74) from *Penicillium islandicum*, destruxin-A4-cyclohydrin (75) from unknown fungi.</figcaption></figure>

For monochlorinated compounds in this cluster, while spontaneous hydrohalogenation involving hydrohalic acid (HCl, HBr) [177] on a double bond can still be considered as an explanation for their formation, we can assume similar halogenation mechanisms inside a family of compounds.

To finish, as already shown, halogenation of alkanes is well represented in fungi. NHFe halogenases and SAM-dependant halide methyltransferases (like AoiQ) are the only known enzymes that can halogenate unactivated aliphatic carbons. However, they are poorly described in fungi. And yet, many fungal compounds carry halogen on aliphatic chains as evidenced in clusters 8, 9 and 10.

Cluster 8 is composed of curtachalasin L (76), I (77) and K (78) together with xylarichalasin A (79) produced by *Xylaria* sp. [178,179] (Figure 32). All these metabolites carry halogenations on cyclohexane rings. Interestingly, during 12-epi-fischerindole G formation catalyzed by the bacterial NHFe halogenases WelO5 in *Hapalosiphon welwischii*, halogenation of a cyclohexane has been described [138]. We can then hypothesize that a similar protein is produced by this fungus and involved in the biosynthesis of these compounds. This hypothesis can be applied for metabolites in cluster 9 where halogenation again occurs on the cyclohexane ring of the three metabolites pestalotiopens A (80), B (81) and C (82) produced by *Pestalotiopsis* sp. [180] (Figure 32). Our search for WelO5 homologous proteins in Xylariales genomes available on MycoCosm gave no result. However, it is possible that even if a fungal NHFe halogenase was present, its sequence might be too different from bacterial NHFe halogenase WelO5.

Cluster 10 is composed of the ribosomally synthesized and post-translationally modified peptides (RiPPs) victorins B (83), D (84) and E (85) from *Cochliobolus victoriae* [181–183] (Figure 33). All these molecules present terminal chlorines on aliphatic chains. The chlorinated lipopeptide barbamide described in the literature from the marine cyanobacterium *Lyngbya majuscula* present the same trichlorinated carbon sp³ as 84. Biosynthesis of barbamide has been investigated and was shown to be performed by a hybrid NRPS/PKS enzyme. In the corresponding biosynthetic cluster, barB1 and barB2 encode two proteins that share homology with NHFe halogenases SyrB1-SyrB2 and CmA B from *Pseudomonas syringae*. These enzymes have been described to halogenate the amino-acid L-threonine for SyrB2 and L-alloisoleucine for CmA B before their insertion into non ribosomal peptides (NRP) [139,184]. For barbamide, trichlorination of leucine was then shown to be catalyzed
by BarB2, when leucine is bound to BarB1 following the NHFe halogenase radical mechanism [184]. In the case of **83**, **84** and **85**, halogenation could be due to the presence of a protein belonging to this same enzymatic family. Indeed, halogenation also occurs on a leucine residue, and we can hypothesize a similar reaction on the amino acid before its insertion in the different peptides. However, ribosomes are described to insert only canonical amino acids in peptides making this hypothesis questionable. Kessler et al. who have worked on victorin biosynthesis hypothesized that NHFe halogenases could be involved but no homolog enzyme is found in the genome of the fungi *C. victoriae*. In fact, difference between fungal and bacterial NHFe halogenases could be important [183].

<figure><img src="image_35.png" /><figcaption>**Figure 32.** Compounds belonging to cluster 8: curtachalasin derivatives L (76), I (77), and K (78) and xylarichalasin A (79); and cluster 9: pestalotiopsins A (80), B (81), and C (82).</figcaption></figure>

<figure><img src="image_36.png" /><figcaption>**Figure 33.** Cluster 10: victorins B (83), D (84), and E (85) from *Cochliobolus victoriae* and bacterial compound barbamide.</figcaption></figure>
by BarB2, when leucine is bound to BarB1 following the NHFe halogenase radical mechanism [184]. In the case of **83**, **84** and **85**, halogenation could be due to the presence of a protein belonging to this same enzymatic family. Indeed, halogenation also occurs on a leucine residue, and we can hypothesize a similar reaction on the amino acid before its insertion in the different peptides. However, ribosomes are described to insert only canonical amino acids in peptides making this hypothesis questionable. Kessler et al. who have worked on victorin biosynthesis hypothesized that NHFe halogenases could be involved but no homolog enzyme is found in the genome of the fungi *C. victoriae*. In fact, difference between fungal and bacterial NHFe halogenases could be important [183].

<figure><img src="image_35.png" /><figcaption>**Figure 32.** Compounds belonging to cluster 8: curtachalasin derivatives L (76), I (77), and K (78) and xylarichalasin A (79); and cluster 9: pestalotiopsins A (80), B (81), and C (82).</figcaption></figure>

<figure><img src="image_36.png" /><figcaption>**Figure 33.** Cluster 10: victorins B (83), D (84), and E (85) from *Cochliobolus victoriae* and bacterial compound barbamide.</figcaption></figure>
## 4. Conclusions

Fungi are great factories of specialised metabolites which can be promoted in many fields such as therapeutics, food industry, agronomy and cosmetics. Fungal secondary metabolism has been widely studied over the last decades and has led to the characterization of numerous enzymatic families, and in the case of this review, of different families of halogenating enzymes. Some are easier to characterize than others because of their specific position in the fungal genome in BGCs such as most of the FDHs. However, for others such as haloperoxidases for example, their characterization remains poor. All classes of halogenating enzymes discovered so far allow to explain much of the wide diversity of halogenated metabolites found in fungal metabolomes. Nevertheless, as reviewed, not many halogenated compounds reported from fungi have had their biosynthesis and more particularly their halogenation mechanism solved. Much remains to be discovered in this field. Regarding the high diversity of halogenated metabolites found in fungi, the diversity of skeletons on which halogen atoms are incorporated also revealed to be beyond the knowledge we have on fungal halogenating enzymes so far.

**Supplementary Materials:** The following supporting information can be downloaded at: https://www.mdpi.com/article/10.3390/molecules27103157/s1, Figure S1: MCD assay: monochlorodimedon is halogenated to obtain dichlorodimedone if X⁺ = Cl⁺ or bromochlorodimedon if X⁺ = Br⁺. Figure S2: Alignment between vHPOs (fungi, algae and bacteria) and bacterial acid phosphatases PAP2. Figure S3: Summary of all putative sequences of fungal halogenation enzymes detected in both UniProt and MycoCosm databases. Distribution of all the detected sequences is found in each pie chart. The number of genomes for each phylogenetic branch available on MycoCosm database are given in grey under each fungal division. Number of sequences found in each database in January 2022 is indicated next to the caption in the summary part. All described enzymes are presented in the rectangle boxes in front of the corresponding fungal division.

**Author Contributions:** Conceptualization, C.R. and L.M.-C.; methodology, B.C., C.R. and L.M.-C.; software, B.C., C.R. and L.M.-C.; validation, C.R., L.M.-C. and Y.F.P.; investigation, B.C.; resources, C.R., L.M.-C., Y.F.P. and O.G.; project administration, C.R., L.M.-C., Y.F.P. and O.G.; funding acquisition, C.R. and L.M.-C.; writing—original draft preparation, B.C.; writing—review and editing, B.C., C.R., L.M.-C., Y.F.P. and O.G.; supervision, C.R., L.M.-C. and Y.F.P. All authors have read and agreed to the published version of the manuscript.

**Funding:** This research was funded by grants from the Ministère de l'Enseignement Supérieur et de la Recherche (Paris, France) and IBSAM (Institut Brestois Santé Agro Matière). This work has been supported by the French National Research Agency (ANR) through the JCJC funding instrument for the "HALO-CAT" project with the reference number ANR-21-CE44-0003-01.

**Institutional Review Board Statement:** Not applicable.

**Informed Consent Statement:** Not applicable.

**Data Availability Statement:** Not applicable.

**Conflicts of Interest:** The authors declare no conflict of interest.

## References

1. Petty, M.A. An Introduction to the Origin and Biochemistry of Microbial Halometabolites. *Bacteriol. Rev.* **1961**, *25*, 111–130.
[CrossRef] [PubMed]

2. Zopf, W. Zur Kenntniss Der Flechtenstoffe (12. Mittheilung.). *Justus Liebigs Ann. Chem.* **1904**, *336*, 46–85. [CrossRef]

3. Morris, D.R.; Hager, L.P. Chloroperoxidase. I. Isolation and Properties of the Crystalline Glycoprotein. *J. Biol. Chem.* **1966**, *241*, 1763–1768. [CrossRef]

4. CRC Press Dictionary of Natural Products (DNP). Available online: https://dnp.chemnetbase.com/ (accessed on 1 March 2021).

5. Gribble, G.W. *Naturally Occuring Organohalogen Compounds—A Comprehensive Update*; Gribble, G.W., Ed.; Springer: Vienna, Austria, 2010; ISBN 978-3-211-99322-4.

6. Dickson, A.G.; Goyet, C. Handbook of Methods for the Analysis of the Various Parameters of the Carbon Dioxide System in Sea Water. Version 2. In *DOE*; Dickson, A.G., Goyet, C., Eds.; Oak Ridge National Laboratory: Oak Ridge, TN, USA, 1994; pp. 1–187.
4. Conclusions

Fungi are great factories of specialised metabolites which can be promoted in many
fields such as therapeutics, food industry, agronomy and cosmetics. Fungal secondary
metabolism has been widely studied over the last decades and has led to the characteriza-
tion of numerous enzymatic families, and in the case of this review, of different families of
halogenating enzymes. Some are easier to characterize than others because of their specific
position in the fungal genome in BGCs such as most of the FDHs. However, for others
such as haloperoxidases for example, their characterization remains poor. All classes of
halogenating enzymes discovered so far allow to explain much of the wide diversity of
halogenated metabolites found in fungal metabolomes. Nevertheless, as reviewed, not
many halogenated compounds reported from fungi have had their biosynthesis and more
particularly their halogenation mechanism solved. Much remains to be discovered in this
field. Regarding the high diversity of halogenated metabolites found in fungi, the diversity
of skeletons on which halogen atoms are incorporated also revealed to be beyond the
knowledge we have on fungal halogenating enzymes so far.

**Supplementary Materials:** The following supporting information can be downloaded at: https://www.mdpi.com/article/10.3390/molecules27103157/s1, Figure S1: MCD assay: monochlorodimedon is halogenated to obtain dichlorodimedone if X⁺ = Cl⁺ or bromochlorodimedon if X⁺ = Br⁺. Figure S2: Alignment between vHPOs (fungi, algae and bacteria) and bacterial acid phosphatases PAP2. Figure S3: Summary of all putative sequences of fungal halogenation enzymes detected in both UniProt and MycoCosm databases. Distribution of all the detected sequences is found in each pie chart. The number of genomes for each phylogenetic branch available on MycoCosm database are given in grey under each fungal division. Number of sequences found in each database in January 2022 is indicated next to the caption in the summary part. All described enzymes are presented in the rectangle boxes in front of the corresponding fungal division.

**Author Contributions:** Conceptualization, C.R. and L.M.-C.; methodology, B.C., C.R. and L.M.-C.; software, B.C., C.R. and L.M.-C.; validation, C.R., L.M.-C. and Y.F.P.; investigation, B.C.; resources, C.R., L.M.-C., Y.F.P. and O.G.; project administration, C.R., L.M.-C., Y.F.P. and O.G.; funding acquisition, C.R. and L.M.-C.; writing—original draft preparation, B.C.; writing—review and editing, B.C., C.R., L.M.-C., Y.F.P. and O.G.; supervision, C.R., L.M.-C. and Y.F.P. All authors have read and agreed to the published version of the manuscript.

**Funding:** This research was funded by grants from the Ministère de l'Enseignement Supérieur et de la Recherche (Paris, France) and IBSAM (Institut Brestois Santé Agro Matière). This work has been supported by the French National Research Agency (ANR) through the JCJC funding instrument for the "HALO-CAT" project with the reference number ANR-21-CE44-0003-01.

**Institutional Review Board Statement:** Not applicable.

**Informed Consent Statement:** Not applicable.

**Data Availability Statement:** Not applicable.

**Conflicts of Interest:** The authors declare no conflict of interest.

References

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   [CrossRef] [PubMed]

2. Zopf, W. Zur Kenntniss Der Flechtenstoffe (12. Mittheilung.). *Justus Liebigs Ann. Chem.* **1904**, *336*, 46–85. [CrossRef]

3. Morris, D.R.; Hager, L.P. Chloroperoxidase. I. Isolation and Properties of the Crystalline Glycoprotein. *J. Biol. Chem.* **1966**, *241*, 1763–1768. [CrossRef]

4. CRC Press Dictionary of Natural Products (DNP). Available online: https://dnp.chemnetbase.com/ (accessed on 1 March 2021).

5. Gribble, G.W. *Naturally Occuring Organohalogen Compounds—A Comprehensive Update*; Gribble, G.W., Ed.; Springer: Vienna, Austria, 2010; ISBN 978-3-211-99322-4.

6. Dickson, A.G.; Goyet, C. Handbook of Methods for the Analysis of the Various Parameters of the Carbon Dioxide System in Sea Water. Version 2. In *DOE*; Dickson, A.G., Goyet, C., Eds.; Oak Ridge National Laboratory: Oak Ridge, TN, USA, 1994; pp. 1–187.
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21. Burgaud, G.; Edgcomb, V.; Hassett, B.T.; Kumar, A.; Li, W.; Mara, P.; Peng, X.; Philippe, A.; Phule, P.; Prado, S.; et al. Marine Fungi. In *The Marine Microbiome*; Stal, L.J., Cretoiu, M.S., Eds.; Springer: Cham, Switzerland, 2022; pp. 243–295. ISBN 978-3-030-90382-4.

22. Wang, C.; Lu, H.; Lan, J.; Zaman, K.; Cao, S. A Review: Halogenated Compounds from Marine Fungi. *Molecules* **2021**, *26*, 458. [CrossRef]

23. Grigoriev, I.V.; Nikitin, R.; Haridas, S.; Kuo, A.; Ohm, R.; Otillar, R.; Riley, R.; Salamov, A.; Zhao, X.; Korzeniewski, F.; et al. MycoCosm Portal: Gearing up for 1000 Fungal Genomes. *Nucleic Acids Res.* **2014**, *42*, 699–704. [CrossRef]

24. Altschul, S.F.; Gish, W.; Miller, W.; Myers, E.W.; Lipman, D.J. Basic Local Alignment Search Tool. *J. Mol. Biol.* **1990**, *215*, 403–410. [CrossRef]

25. Kumar, S.; Stecher, G.; Li, M.; Knyaz, C.; Tamura, K. MEGA X: Molecular Evolutionary Genetics Analysis across Computing Platforms. *Mol. Biol. Evol.* **2018**, *35*, 1547–1549. [CrossRef] [PubMed]

26. Sehnal, D.; Rose, A.S.; Koča, J.; Burley, S.K.; Velankar, S. Mol*: Towards a Common Library and Tools for Web Molecular Graphics. In *MolVA '18, Proceedings of the Workshop on Molecular Graphics and Visual Analysis of Molecular Data*; Byska, J., Krone, M., Sommer, B., Eds.; The Eurographics Association: Goslar, Germany, 2018; ISBN 978-3-03868-061-1.

27. Fang, G.-H.; Kenigsberg, P.; Axley, M.J.; Nuell, M.; Hager, L.P. Cloning and Sequencing of Chloroperoxidase CDNA. *Nucleic Acids Res.* **1986**, *14*, 8061–8071. [CrossRef] [PubMed]

28. Simons, B.H.; Barnett, P.; Vollenbroek, E.G.M.; Dekker, H.L.; Muijsers, A.O.; Messerschmidt, A.; Wever, R. Primary Structure and Characterization of the Vanadium Chloroperoxidase from the Fungus Curvularia Inaequalis. *Eur. J. Biochem.* **1995**, *229*, 566–574. [CrossRef] [PubMed]

29. Barnett, P.; Hemrika, W.; Dekker, H.L.; Muijsers, A.O.; Renirie, R.; Wever, R. Isolation, Characterization, and Primary Structure of the Vanadium Chloroperoxidase from the Fungus Embellisia Didymospora. *J. Biol. Chem.* **1998**, *273*, 23381–23387. [CrossRef] [PubMed]

30. Fraley, A.E.; Garcia-borràs, M.; Tripathi, A.; Khare, D.; Eduardo, V.; Tran, H.; Dan, Q.; Webb, G.P.; Watts, K.R.; Crews, P.; et al. Function and Structure of MalA/MalA', Iterative Halogenases for Late-Stage C–H Functionalization of Indole Alkaloids. *J. Am. Chem. Soc.* **2017**, *139*, 12060–12068. [CrossRef]

31. Reeves, C.D.; Hu, Z.; Reid, R.; Kealey, J.T. Genes for the Biosynthesis of the Fungal Polyketides Hypothemycin from Hypomyces Subiculosus and Radicicol from Pochonia Chlamydosporia. *Appl. Environ. Microbiol.* **2008**, *74*, 5121–5129. [CrossRef]
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14. Julianti, E.; Abrian, I.A.; Wibowo, M.S.; Azhari, M.; Tsurayya, N.; Izzati, F.; Juanssilfero, A.B.; Bayu, A.; Rahmawati, S.I.; Putra, M.Y. Secondary Metabolites from Marine-Derived Fungi and Actinobacteria as Potential Sources of Novel Colorectal Cancer Drugs. *Mar. Drugs* **2022**, *20*, 67. [CrossRef] [PubMed]

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19. Ma, X.; Yu, J.; Jiang, M.; Wang, M.; Tang, L.; Wei, M.; Zhou, Q. Mild and Regioselective Bromination of Phenols with TMSBr. *Eur. J. Org. Chem.* **2019**, *2019*, 4593–4596. [CrossRef]

20. Tang, R.-J.; Milcent, T.; Crousse, B. Regioselective Halogenation of Arenes and Heterocycles in Hexafluoroisopropanol. *J. Org. Chem.* **2018**, *83*, 930–938. [CrossRef]

21. Burgaud, G.; Edgcomb, V.; Hassett, B.T.; Kumar, A.; Li, W.; Mara, P.; Peng, X.; Philippe, A.; Phule, P.; Prado, S.; et al. Marine Fungi. In *The Marine Microbiome*; Stal, L.J., Cretoiu, M.S., Eds.; Springer: Cham, Switzerland, 2022; pp. 243–295. ISBN 978-3-030-90382-4.

22. Wang, C.; Lu, H.; Lan, J.; Zaman, K.; Cao, S. A Review: Halogenated Compounds from Marine Fungi. *Molecules* **2021**, *26*, 458. [CrossRef]

23. Grigoriev, I.V.; Nikitin, R.; Haridas, S.; Kuo, A.; Ohm, R.; Otillar, R.; Riley, R.; Salamov, A.; Zhao, X.; Korzeniewski, F.; et al. MycoCosm Portal: Gearing up for 1000 Fungal Genomes. *Nucleic Acids Res.* **2014**, *42*, 699–704. [CrossRef]

24. Altschul, S.F.; Gish, W.; Miller, W.; Myers, E.W.; Lipman, D.J. Basic Local Alignment Search Tool. *J. Mol. Biol.* **1990**, *215*, 403–410. [CrossRef]

25. Kumar, S.; Stecher, G.; Li, M.; Knyaz, C.; Tamura, K. MEGA X: Molecular Evolutionary Genetics Analysis across Computing Platforms. *Mol. Biol. Evol.* **2018**, *35*, 1547–1549. [CrossRef] [PubMed]

26. Sehnal, D.; Rose, A.S.; Koča, J.; Burley, S.K.; Velankar, S. Mol*: Towards a Common Library and Tools for Web Molecular Graphics. In *MolVA '18, Proceedings of the Workshop on Molecular Graphics and Visual Analysis of Molecular Data*; Byska, J., Krone, M., Sommer, B., Eds.; The Eurographics Association: Goslar, Germany, 2018; ISBN 978-3-03868-061-1.

27. Fang, G.-H.; Kenigsberg, P.; Axley, M.J.; Nuell, M.; Hager, L.P. Cloning and Sequencing of Chloroperoxidase CDNA. *Nucleic Acids Res.* **1986**, *14*, 8061–8071. [CrossRef] [PubMed]

28. Simons, B.H.; Barnett, P.; Vollenbroek, E.G.M.; Dekker, H.L.; Muijsers, A.O.; Messerschmidt, A.; Wever, R. Primary Structure and Characterization of the Vanadium Chloroperoxidase from the Fungus Curvularia Inaequalis. *Eur. J. Biochem.* **1995**, *229*, 566–574. [CrossRef] [PubMed]

29. Barnett, P.; Hemrika, W.; Dekker, H.L.; Muijsers, A.O.; Renirie, R.; Wever, R. Isolation, Characterization, and Primary Structure of the Vanadium Chloroperoxidase from the Fungus Embellisia Didymospora. *J. Biol. Chem.* **1998**, *273*, 23381–23387. [CrossRef] [PubMed]

30. Fraley, A.E.; Garcia-borràs, M.; Tripathi, A.; Khare, D.; Eduardo, V.; Tran, H.; Dan, Q.; Webb, G.P.; Watts, K.R.; Crews, P.; et al. Function and Structure of MalA/MalA', Iterative Halogenases for Late-Stage C–H Functionalization of Indole Alkaloids. *J. Am. Chem. Soc.* **2017**, *139*, 12060–12068. [CrossRef]

31. Reeves, C.D.; Hu, Z.; Reid, R.; Kealey, J.T. Genes for the Biosynthesis of the Fungal Polyketides Hypothemycin from Hypomyces Subiculosus and Radicicol from Pochonia Chlamydosporia. *Appl. Environ. Microbiol.* **2008**, *74*, 5121–5129. [CrossRef]
32. Wang, S.; Xu, Y.; Maine, E.A.; Wijeratne, E.M.K.; Espinosa-Artiles, P.; Gunatilaka, A.A.L.; Molnár, I. Functional Characterization of the Biosynthesis of Radicicol, an Hsp90 Inhibitor Resorcylic Acid Lactone from Chaetomium Chiversii. *Chem. Biol.* **2008**, *15*, 1328–1338. [CrossRef]

33. Ferrara, M.; Perrone, G.; Gambacorta, L.; Epifani, F.; Solfrizzo, M.; Gallo, A. Identification of a Halogenase Involved in the Biosynthesis of Ochratoxin A in Aspergillus Carbonarius. *Appl. Environ. Microbiol.* **2016**, *82*, 5631–5641. [CrossRef]

34. Susca, A.; Proctor, R.H.; Morelli, M.; Haidukowski, M.; Gallo, A.; Logrieco, A.F.; Moretti, A. Variation in Fumonisin and Ochratoxin Production Associated with Differences in Biosynthetic Gene Content in Aspergillus Niger and A. Welwitschiae Isolates from Multiple Crop and Geographic Origins. *Front. Microbiol.* **2016**, *7*, 1–15. [CrossRef]

35. Xu, X.; Liu, L.; Zhang, F.; Wang, W.; Li, J.; Guo, L.; Che, Y.; Liu, G. Identification of the First Diphenyl Ether Gene Cluster for Pestheic Acid Biosynthesis in Plant Endophyte Pestalotiopsis Fici. *ChemBioChem* **2014**, *15*, 284–292. [CrossRef]

36. Nielsen, M.T.; Nielsen, J.B.; Anyaogu, D.C.; Holm, D.K.; Nielsen, K.F.; Larsen, T.O.; Mortensen, U.H. Heterologous Reconstitution of the Intact Geodin Gene Cluster in Aspergillus Nidulans through a Simple and Versatile PCR Based Approach. *PLoS ONE* **2013**, *8*, e72871. [CrossRef]

37. Cacho, R.A.; Chooi, Y.H.; Zhou, H.; Tang, Y. Complexity Generation in Fungal Polyketide Biosynthesis: A Spirocycle-Forming P450 in the Concise Pathway to the Antifungal Drug Griseofulvin. *ACS Chem. Biol.* **2013**, *8*, 2322–2330. [CrossRef] [PubMed]

38. Chankhamjon, P.; Boettger-Schmidt, D.; Scherlach, K.; Urbansky, B.; Lackner, G.; Kalb, D.; Dahse, H.M.; Hoffmeister, D.; Hertweck, C. Biosynthesis of the Halogenated Mycotoxin Aspirochlorine in Koji Mold Involves a Cryptic Amino Acid Conversion. *Angew. Chem.-Int. Ed.* **2014**, *53*, 13409–13413. [CrossRef] [PubMed]

39. Araki, Y.; Awakawa, T.; Matsuzaki, M.; Cho, R.; Matsuda, Y.; Hoshino, S.; Shinohara, Y.; Yamamoto, M.; Kido, Y.; Inaoka, D.K.; et al. Complete Biosynthetic Pathways of Ascofuranone and Ascochlorin in Acremonium Egyptiacum. *Proc. Natl. Acad. Sci. USA* **2019**, *116*, 8269–8274. [CrossRef]

40. Wick, J.; Heine, D.; Lackner, G.; Misiek, M.; Tauber, J.; Jagusch, H.; Hertweck, C.; Hoffmeister, D. A Fivefold Parallelized Biosynthetic Process Secures Chlorination of Armillaria Mellea (Honey Mushroom) Toxins. *Appl. Environ. Microbiol.* **2016**, *82*, 1196–1204. [CrossRef]

41. Ugai, T.; Minami, A.; Tanaka, S.; Ozaki, T.; Liu, C.; Shigemori, H.; Hashimoto, M.; Oikawa, H. Biosynthetic Machinery of 6-Hydroxymellein Derivatives Leading to Cyclohelminthols and Palmaenones. *ChemBioChem* **2020**, *21*, 360–367. [CrossRef]

42. Winter, J.M.; Sato, M.; Sugimoto, S.; Chiou, G.; Garg, N.K.; Tang, Y.; Watanabe, K. Identification and Characterization of the Chaetoviridin and Chaetomugilin Gene Cluster in Chaetomium Globosum Reveal Dual Functions of an Iterative Highly-Reducing Polyketide Synthase. *J. Am. Chem. Soc.* **2012**, *134*, 17900–17903. [CrossRef]

43. Axley, M.J.; Kenigsberg, P.; Hager, L.P. Fructose Induces and Glucose Represses Chloroperoxidase mRNA Levels. *J. Biol. Chem.* **1986**, *261*, 15058–15061. [CrossRef]

44. Blin, K.; Shaw, S.; Kloosterman, A.M.; Charlop-Powers, Z.; van Wezel, G.P.; Medema, M.H.; Weber, T. AntiSMASH 6.0: Improving Cluster Detection and Comparison Capabilities. *Nucleic Acids Res.* **2021**, *49*, W29–W35. [CrossRef]

45. Menon, B.R.K.; Richmond, D.; Menon, N. Halogenases for Biosynthetic Pathway Engineering: Toward New Routes to Naturals and Non-Naturals. *Catal. Rev.* **2020**, *62*, 1–59. [CrossRef]

46. Crowe, C.; Molyneux, S.; Sharma, S.V.; Zhang, Y.; Gkotsi, D.S.; Connaris, H.; Goss, R.J.M. Halogenases: A Palette of Emerging Opportunities for Synthetic Biology–Synthetic Chemistry and C–H Functionalisation. *Chem. Soc. Rev.* **2021**, *50*, 9443–9481. [CrossRef] [PubMed]

47. Dachwitz, S.; Widmann, C.; Frese, M.; Niemann, H.H.; Sewald, N. Enzymatic Halogenation: Enzyme Mining, Mechanisms, and Implementation in Reaction Cascades. In *Amino Acids, Peptides and Proteins*; Royal Society of Chemistry: Cambridge, UK, 2020; Volume 44, pp. 1–43.

48. Hager, L.; Morris, D.; Brown, F.; Eberwein, H. Chloroperoxidase: Utilization of Halogen Anions. *J. Biol. Chem.* **1966**, *241*, 1769–1777. [CrossRef]

49. Verhaeghe, E.; Buisson, D.; Zekri, E.; Leblanc, C.; Potin, P.; Ambroise, Y. A Colorimetric Assay for Steady-State Analyses of Iodo- and Bromoperoxidase Activities. *Anal. Biochem.* **2008**, *379*, 60–65. [CrossRef] [PubMed]

50. Jordan, P.; Vilter, H. Native Bromoperoxidases Do Not Bind to Nitrocellulose: Use of DEAE-Cellulose as an Alternative in Blotting. *Electrophoresis* **1990**, *11*, 653–655. [CrossRef] [PubMed]

51. Nag, N.; Khan, H.; Tripathi, T. Strategies to Improve the Expression and Solubility of Recombinant Proteins in *E. Coli*. In *Advances in Protein Molecular and Structural Biology Methods*; Elsevier: Amsterdam, The Netherlands, 2022; pp. 1–12.

52. Zemella, A.; Thoring, L.; Hoffmeister, C.; Kubick, S. Cell-Free Protein Synthesis: Pros and Cons of Prokaryotic and Eukaryotic Systems. *ChemBioChem* **2015**, *16*, 2420–2431. [CrossRef]

53. Mamipour, M.; Yousefi, M.; Hasanzadeh, M. An Overview on Molecular Chaperones Enhancing Solubility of Expressed Recombinant Proteins with Correct Folding. *Int. J. Biol. Macromol.* **2017**, *102*, 367–375. [CrossRef] [PubMed]

54. Hemrika, W.; Renirie, R.; Macedo-Ribeiro, S.; Messerschmidt, A.; Wever, R. Heterologous Expression of the Vanadium-Containing Chloroperoxidase from Curvularia Inaequalis in Saccharomyces Cerevisiae and Site-Directed Mutagenesis of the Active Site Residues His496, Lys353, Arg360, and Arg490. *J. Biol. Chem.* **1999**, *274*, 23820–23827. [CrossRef]

55. Conesa, A.; van de Velde, F.; van Rantwijk, F.; Sheldon, R.A.; van den Hondel, C.A.M.J.J.; Punt, P.J. Expression of the Caldariomyces Fumago Chloroperoxidase in Aspergillus Niger and Characterization of the Recombinant Enzyme. *J. Biol. Chem.* **2001**, *276*, 17635–17640. [CrossRef]
32. Wang, S.; Xu, Y.; Maine, E.A.; Wijeratne, E.M.K.; Espinosa-Artiles, P.; Gunatilaka, A.A.L.; Molnár, I. Functional Characterization of the Biosynthesis of Radicicol, an Hsp90 Inhibitor Resorcylic Acid Lactone from Chaetomium Chiversii. *Chem. Biol.* **2008**, *15*, 1328–1338. [CrossRef]

33. Ferrara, M.; Perrone, G.; Gambacorta, L.; Epifani, F.; Solfrizzo, M.; Gallo, A. Identification of a Halogenase Involved in the Biosynthesis of Ochratoxin A in Aspergillus Carbonarius. *Appl. Environ. Microbiol.* **2016**, *82*, 5631–5641. [CrossRef]

34. Susca, A.; Proctor, R.H.; Morelli, M.; Haidukowski, M.; Gallo, A.; Logrieco, A.F.; Moretti, A. Variation in Fumonisin and Ochratoxin Production Associated with Differences in Biosynthetic Gene Content in Aspergillus Niger and A. Welwitschiae Isolates from Multiple Crop and Geographic Origins. *Front. Microbiol.* **2016**, *7*, 1–15. [CrossRef]

35. Xu, X.; Liu, L.; Zhang, F.; Wang, W.; Li, J.; Guo, L.; Che, Y.; Liu, G. Identification of the First Diphenyl Ether Gene Cluster for Pestheic Acid Biosynthesis in Plant Endophyte Pestalotiopsis Fici. *ChemBioChem* **2014**, *15*, 284–292. [CrossRef]

36. Nielsen, M.T.; Nielsen, J.B.; Anyaogu, D.C.; Holm, D.K.; Nielsen, K.F.; Larsen, T.O.; Mortensen, U.H. Heterologous Reconstitution of the Intact Geodin Gene Cluster in Aspergillus Nidulans through a Simple and Versatile PCR Based Approach. *PLoS ONE* **2013**, *8*, e72871. [CrossRef]

37. Cacho, R.A.; Chooi, Y.H.; Zhou, H.; Tang, Y. Complexity Generation in Fungal Polyketide Biosynthesis: A Spirocycle-Forming P450 in the Concise Pathway to the Antifungal Drug Griseofulvin. *ACS Chem. Biol.* **2013**, *8*, 2322–2330. [CrossRef] [PubMed]

38. Chankhamjon, P.; Boettger-Schmidt, D.; Scherlach, K.; Urbansky, B.; Lackner, G.; Kalb, D.; Dahse, H.M.; Hoffmeister, D.; Hertweck, C. Biosynthesis of the Halogenated Mycotoxin Aspirochlorine in Koji Mold Involves a Cryptic Amino Acid Conversion. *Angew. Chem.-Int. Ed.* **2014**, *53*, 13409–13413. [CrossRef] [PubMed]

39. Araki, Y.; Awakawa, T.; Matsuzaki, M.; Cho, R.; Matsuda, Y.; Hoshino, S.; Shinohara, Y.; Yamamoto, M.; Kido, Y.; Inaoka, D.K.; et al. Complete Biosynthetic Pathways of Ascofuranone and Ascochlorin in Acremonium Egyptiacum. *Proc. Natl. Acad. Sci. USA* **2019**, *116*, 8269–8274. [CrossRef]

40. Wick, J.; Heine, D.; Lackner, G.; Misiek, M.; Tauber, J.; Jagusch, H.; Hertweck, C.; Hoffmeister, D. A Fivefold Parallelized Biosynthetic Process Secures Chlorination of Armillaria Mellea (Honey Mushroom) Toxins. *Appl. Environ. Microbiol.* **2016**, *82*, 1196–1204. [CrossRef]

41. Ugai, T.; Minami, A.; Tanaka, S.; Ozaki, T.; Liu, C.; Shigemori, H.; Hashimoto, M.; Oikawa, H. Biosynthetic Machinery of 6-Hydroxymellein Derivatives Leading to Cyclohelminthols and Palmaenones. *ChemBioChem* **2020**, *21*, 360–367. [CrossRef]

42. Winter, J.M.; Sato, M.; Sugimoto, S.; Chiou, G.; Garg, N.K.; Tang, Y.; Watanabe, K. Identification and Characterization of the Chaetoviridin and Chaetomugilin Gene Cluster in Chaetomium Globosum Reveal Dual Functions of an Iterative Highly-Reducing Polyketide Synthase. *J. Am. Chem. Soc.* **2012**, *134*, 17900–17903. [CrossRef]

43. Axley, M.J.; Kenigsberg, P.; Hager, L.P. Fructose Induces and Glucose Represses Chloroperoxidase mRNA Levels. *J. Biol. Chem.* **1986**, *261*, 15058–15061. [CrossRef]

44. Blin, K.; Shaw, S.; Kloosterman, A.M.; Charlop-Powers, Z.; van Wezel, G.P.; Medema, M.H.; Weber, T. AntiSMASH 6.0: Improving Cluster Detection and Comparison Capabilities. *Nucleic Acids Res.* **2021**, *49*, W29–W35. [CrossRef]

45. Menon, B.R.K.; Richmond, D.; Menon, N. Halogenases for Biosynthetic Pathway Engineering: Toward New Routes to Naturals and Non-Naturals. *Catal. Rev.* **2020**, *62*, 1–59. [CrossRef]

46. Crowe, C.; Molyneux, S.; Sharma, S.V.; Zhang, Y.; Gkotsi, D.S.; Connaris, H.; Goss, R.J.M. Halogenases: A Palette of Emerging Opportunities for Synthetic Biology–Synthetic Chemistry and C–H Functionalisation. *Chem. Soc. Rev.* **2021**, *50*, 9443–9481. [CrossRef] [PubMed]

47. Dachwitz, S.; Widmann, C.; Frese, M.; Niemann, H.H.; Sewald, N. Enzymatic Halogenation: Enzyme Mining, Mechanisms, and Implementation in Reaction Cascades. In *Amino Acids, Peptides and Proteins*; Royal Society of Chemistry: Cambridge, UK, 2020; Volume 44, pp. 1–43.

48. Hager, L.; Morris, D.; Brown, F.; Eberwein, H. Chloroperoxidase: Utilization of Halogen Anions. *J. Biol. Chem.* **1966**, *241*, 1769–1777. [CrossRef]

49. Verhaeghe, E.; Buisson, D.; Zekri, E.; Leblanc, C.; Potin, P.; Ambroise, Y. A Colorimetric Assay for Steady-State Analyses of Iodo- and Bromoperoxidase Activities. *Anal. Biochem.* **2008**, *379*, 60–65. [CrossRef] [PubMed]

50. Jordan, P.; Vilter, H. Native Bromoperoxidases Do Not Bind to Nitrocellulose: Use of DEAE-Cellulose as an Alternative in Blotting. *Electrophoresis* **1990**, *11*, 653–655. [CrossRef] [PubMed]

51. Nag, N.; Khan, H.; Tripathi, T. Strategies to Improve the Expression and Solubility of Recombinant Proteins in *E. Coli*. In *Advances in Protein Molecular and Structural Biology Methods*; Elsevier: Amsterdam, The Netherlands, 2022; pp. 1–12.

52. Zemella, A.; Thoring, L.; Hoffmeister, C.; Kubick, S. Cell-Free Protein Synthesis: Pros and Cons of Prokaryotic and Eukaryotic Systems. *ChemBioChem* **2015**, *16*, 2420–2431. [CrossRef]

53. Mamipour, M.; Yousefi, M.; Hasanzadeh, M. An Overview on Molecular Chaperones Enhancing Solubility of Expressed Recombinant Proteins with Correct Folding. *Int. J. Biol. Macromol.* **2017**, *102*, 367–375. [CrossRef] [PubMed]

54. Hemrika, W.; Renirie, R.; Macedo-Ribeiro, S.; Messerschmidt, A.; Wever, R. Heterologous Expression of the Vanadium-Containing Chloroperoxidase from Curvularia Inaequalis in Saccharomyces Cerevisiae and Site-Directed Mutagenesis of the Active Site Residues His496, Lys353, Arg360, and Arg490. *J. Biol. Chem.* **1999**, *274*, 23820–23827. [CrossRef]

55. Conesa, A.; van de Velde, F.; van Rantwijk, F.; Sheldon, R.A.; van den Hondel, C.A.M.J.J.; Punt, P.J. Expression of the Caldariomyces Fumago Chloroperoxidase in Aspergillus Niger and Characterization of the Recombinant Enzyme. *J. Biol. Chem.* **2001**, *276*, 17635–17640. [CrossRef]
56. Zhou, H.; Qiao, K.; Gao, Z.; Vederas, J.C.; Tang, Y. Insights into Radicicol Biosynthesis via Heterologous Synthesis of Intermediates and Analogs. *J. Biol. Chem.* **2010**, *285*, 41412–41421. [CrossRef]

57. Messerschmidt, A.; Prade, L.; Wever, R. Implications for the Catalytic Mechanism of the Vanadium-Containing Enzyme Chloroperoxidase from the Fungus Curvularia Inaequalis by X-Ray Structures of the Native and Peroxide Form. *Biol. Chem.* **1997**, *378*, 309–316. [CrossRef]

58. Pickard, M.A.; Kadima, T.A.; Carmichael, R.D. Chloroperoxidase, a Peroxidase with Potential. *J. Ind. Microbiol.* **1991**, *7*, 235–241. [CrossRef]

59. Bormann, S.; Gomez Baraibar, A.; Ni, Y.; Holtmann, D.; Hollmann, F. Specific Oxyfunctionalisations Catalysed by Peroxygenases: Opportunities, Challenges and Solutions. *Catal. Sci. Technol.* **2015**, *5*, 2038–2052. [CrossRef]

60. Aoun, S.; Baboulène, M. Regioselective Bromohydroxylation of Alkenes Catalyzed by Chloroperoxidase: Advantages of the Immobilization of Enzyme on Talc. *J. Mol. Catal.-B Enzym.* **1998**, *4*, 101–109. [CrossRef]

61. Wischang, D.; Hartung, J. Bromination of Phenols in Bromoperoxidase-Catalyzed Oxidations. *Tetrahedron* **2012**, *68*, 9456–9463. [CrossRef]

62. van Deurzen, M.P.J.; van Rantwijk, F.; Sheldon, R.A. Synthesis of Substituted Oxindoles by Chloroperoxidase Catalyzed Oxidation of Indoles. *J. Mol. Catal. B: Enzym.* **1996**, *2*, 33–42. [CrossRef]

63. Höfler, G.T.; But, A.; Hollmann, F. Haloperoxidases as Catalysts in Organic Synthesis. *Org. Biomol. Chem.* **2019**, *17*, 9267–9274. [CrossRef]

64. Fernández-Fueyo, E.; H Younes, S.H.; van Rootselaar, S.; Aben, W.M.; Renirie, R.; Wever, R.; Holtmann, D.; J T Rutjes, F.P.; Hollmann, F. A Biocatalytic Aza-Achmatowicz Reaction. *ACS Catal.* **2016**, *6*, 5904–5907. [CrossRef]

65. Yaipakdee, P.; Robertson, L.W. Enzymatic Halogenation of Flavanones and Flavones. *Phytochemistry* **2001**, *57*, 341–347. [CrossRef]

66. Franssen, M.C.R.; van der Plas, H.C. The Chlorination of Barbituric Acid and Some of Its Derivatives by Chloroperoxidase. *Bioorg. Chem.* **1987**, *15*, 59–70. [CrossRef]

67. Geigert, J.; Neidleman, S.L.; Dalietos, D.J. Novel Haloperoxidase Substrates. *Alkynes and Cyclopropanes. J. Biol. Chem.* **1983**, *258*, 2273–2277. [CrossRef]

68. Vázquez-Duhalt, R.; Ayala, M.; Márquez-Rocha, F.J. Biocatalytic Chlorination of Aromatic Hydrocarbons by Chloroperoxidase of Caldariomyces Fumago. *Phytochemistry* **2001**, *58*, 929–933. [CrossRef]

69. Younes, S.H.H.; Tieves, F.; Lan, D.; Wang, Y.; Süss, P.; Brundiek, H.; Wever, R.; Hollmann, F. Chemoenzymatic Halocyclization of γ,δ-Unsaturated Carboxylic Acids and Alcohols. *ChemSusChem* **2019**, *12*, 1–6. [CrossRef]

70. Zeng, J.; Zhan, J. A Novel Fungal Flavin-Dependent Halogenase for Natural Product Biosynthesis. *ChemBioChem* **2010**, *11*, 2119–2123. [CrossRef] [PubMed]

71. Zeng, J.; Lytle, A.K.; Gage, D.; Johnson, S.J.; Zhan, J. Specific Chlorination of Isoquinolines by a Fungal Flavin-Dependent Halogenase. *Bioorg. Med. Chem. Lett.* **2013**, *23*, 1001–1003. [CrossRef]

72. Timmins, A.; De Visser, S.P. Enzymatic Halogenases and Haloperoxidases: Computational Studies on Mechanism and Function. *Adv. Protein Chem. Struct. Biol.* **2015**, *100*, 113–151. [CrossRef]

73. Sundaramoorthy, M.; Terner, J.; Poulos, T.L. The Crystal Structure of Chloroperoxidase: A Heme Peroxidase-Cytochrome P450 Functional Hybrid. *Structure* **1995**, *3*, 1367–1378. [CrossRef]

74. Zong, Q.; Osmulski, P.A.; Hager, L.P. High-Pressure-Assisted Reconstitution of Recombinant Chloroperoxidase. *Biochemistry* **1995**, *34*, 12420–12425. [CrossRef]

75. Butler, A.; Sandy, M. Mechanistic Considerations of Halogenating Enzymes. *Nature* **2009**, *460*, 848–854. [CrossRef]

76. Kühnel, K.; Blankenfeldt, W.; Terner, J.; Schlichting, I. Crystal Structures of Chloroperoxidase with Its Bound Substrates and Complexed with Formate, Acetate, and Nitrate. *J. Biol. Chem.* **2006**, *281*, 23990–23998. [CrossRef]

77. Sundaramoorthy, M.; Terner, J.; Poulos, T.L. Stereochemistry of the Chloroperoxidase Active Site: Crystallographic and Molecular-Modeling Studies. *Chem. Biol.* **1998**, *5*, 461–473. [CrossRef]

78. Shaik, S.; Kumar, D.; De Visser, S.P. A Valence Bond Modeling of Trends in Hydrogen Abstraction Barriers and Transition States of Hydroxylation Reactions Catalyzed by Cytochrome P450 Enzymes. *J. Am. Chem. Soc.* **2008**, *130*, 10128–10140. [CrossRef] [PubMed]

79. Hofrichter, M.; Ullrich, R. Heme-Thiolate Haloperoxidases: Versatile Biocatalysts with Biotechnological and Environmental Significance. *Appl. Microbiol. Biotechnol.* **2006**, *71*, 276–288. [CrossRef] [PubMed]

80. Farhangrazi, Z.S.; Sinclair, R.; Yamazaki, I.; Powers, L.S. Haloperoxidase Activity of Phanerochaete Chrysosporium Lignin Peroxidases H2 and H8. *Biochemistry* **1992**, *31*, 10763–10768. [CrossRef]

81. Ullrich, R.; Nueske, J.; Scheibner, K.; Spantzel, J.; Hofrichter, M. Novel Haloperoxidase from the Agaric Basidiomycete. *Appl. Environ. Microbiol.* **2004**, *70*, 4575–4581. [CrossRef]

82. Piontek, K.; Ullrich, R.; Liers, C.; Diederichs, K.; Plattner, D.A.; Hofrichter, M. Crystallization of a 45 KDa Peroxynase/Peroxidase from the Mushroom Agrocybe Aegerita and Structure Determination by SAD Utilizing Only the Haem Iron. *Acta Crystallogr. Sect. F: Struct. Biol. Cryst. Commun.* **2010**, *66*, 693–698. [CrossRef] [PubMed]

83. Zamocky, M.; Obinger, C. *Biocatalysis Based on Heme Peroxidases*; Torres, E., Ayala, M., Eds.; Springer: Berlin/Heidelberg, Germany, 2010; ISBN 978-3-642-12626-0.

84. China, H.; Okada, Y.; Dohi, T. The Multiple Reactions in the Monochlorodimedone Assay: Discovery of Unique Dehalolactonizations under Mild Conditions. *Asian J. Org. Chem.* **2015**, *4*, 1065–1074. [CrossRef]
56. Zhou, H.; Qiao, K.; Gao, Z.; Vederas, J.C.; Tang, Y. Insights into Radicicol Biosynthesis via Heterologous Synthesis of Intermediates and Analogs. *J. Biol. Chem.* **2010**, *285*, 41412-41421. [CrossRef]

57. Messerschmidt, A.; Prade, L.; Wever, R. Implications for the Catalytic Mechanism of the Vanadium-Containing Enzyme Chloroperoxidase from the Fungus Curvularia Inaequalis by X-Ray Structures of the Native and Peroxide Form. *Biol. Chem.* **1997**, *378*, 309-316. [CrossRef]

58. Pickard, M.A.; Kadima, T.A.; Carmichael, R.D. Chloroperoxidase, a Peroxidase with Potential. *J. Ind. Microbiol.* **1991**, *7*, 235-241. [CrossRef]

59. Bormann, S.; Gomez Baraibar, A.; Ni, Y.; Holtmann, D.; Hollmann, F. Specific Oxyfunctionalisations Catalysed by Peroxygenases: Opportunities, Challenges and Solutions. *Catal. Sci. Technol.* **2015**, *5*, 2038-2052. [CrossRef]

60. Aoun, S.; Baboulène, M. Regioselective Bromohydroxylation of Alkenes Catalyzed by Chloroperoxidase: Advantages of the Immobilization of Enzyme on Talc. *J. Mol. Catal.-B Enzym.* **1998**, *4*, 101-109. [CrossRef]

61. Wischang, D.; Hartung, J. Bromination of Phenols in Bromoperoxidase-Catalyzed Oxidations. *Tetrahedron* **2012**, *68*, 9456-9463. [CrossRef]

62. van Deurzen, M.P.J.; van Rantwijk, F.; Sheldon, R.A. Synthesis of Substituted Oxindoles by Chloroperoxidase Catalyzed Oxidation of Indoles. *J. Mol. Catal. B: Enzym.* **1996**, *2*, 33-42. [CrossRef]

63. Höfler, G.T.; But, A.; Hollmann, F. Haloperoxidases as Catalysts in Organic Synthesis. *Org. Biomol. Chem.* **2019**, *17*, 9267-9274. [CrossRef]

64. Fernández-Fueyo, E.; H Younes, S.H.; van Rootselaar, S.; Aben, W.M.; Renirie, R.; Wever, R.; Holtmann, D.; J T Rutjes, F.P.; Hollmann, F. A Biocatalytic Aza-Achmatowicz Reaction. *ACS Catal.* **2016**, *6*, 5904-5907. [CrossRef]

65. Yaipakdee, P.; Robertson, L.W. Enzymatic Halogenation of Flavanones and Flavones. *Phytochemistry* **2001**, *57*, 341-347. [CrossRef]

66. Franssen, M.C.R.; van der Plas, H.C. The Chlorination of Barbituric Acid and Some of Its Derivatives by Chloroperoxidase. *Bioorg. Chem.* **1987**, *15*, 59-70. [CrossRef]

67. Geigert, J.; Neidleman, S.L.; Dalietos, D.J. Novel Haloperoxidase Substrates. *Alkynes and Cyclopropanes. J. Biol. Chem.* **1983**, *258*, 2273-2277. [CrossRef]

68. Vázquez-Duhalt, R.; Ayala, M.; Márquez-Rocha, F.J. Biocatalytic Chlorination of Aromatic Hydrocarbons by Chloroperoxidase of Caldariomyces Fumago. *Phytochemistry* **2001**, *58*, 929-933. [CrossRef]

69. Younes, S.H.H.; Tieves, F.; Lan, D.; Wang, Y.; Süss, P.; Brundiek, H.; Wever, R.; Hollmann, F. Chemoenzymatic Halocyclization of γ,δ-Unsaturated Carboxylic Acids and Alcohols. *ChemSusChem* **2019**, *12*, 1-6. [CrossRef]

70. Zeng, J.; Zhan, J. A Novel Fungal Flavin-Dependent Halogenase for Natural Product Biosynthesis. *ChemBioChem* **2010**, *11*, 2119-2123. [CrossRef] [PubMed]

71. Zeng, J.; Lytle, A.K.; Gage, D.; Johnson, S.J.; Zhan, J. Specific Chlorination of Isoquinolines by a Fungal Flavin-Dependent Halogenase. *Bioorg. Med. Chem. Lett.* **2013**, *23*, 1001-1003. [CrossRef]

72. Timmins, A.; De Visser, S.P. Enzymatic Halogenases and Haloperoxidases: Computational Studies on Mechanism and Function. *Adv. Protein Chem. Struct. Biol.* **2015**, *100*, 113-151. [CrossRef]

73. Sundaramoorthy, M.; Terner, J.; Poulos, T.L. The Crystal Structure of Chloroperoxidase: A Heme Peroxidase-Cytochrome P450 Functional Hybrid. *Structure* **1995**, *3*, 1367-1378. [CrossRef]

74. Zong, Q.; Osmulski, P.A.; Hager, L.P. High-Pressure-Assisted Reconstitution of Recombinant Chloroperoxidase. *Biochemistry* **1995**, *34*, 12420-12425. [CrossRef]

75. Butler, A.; Sandy, M. Mechanistic Considerations of Halogenating Enzymes. *Nature* **2009**, *460*, 848-854. [CrossRef]

76. Kühnel, K.; Blankenfeldt, W.; Terner, J.; Schlichting, I. Crystal Structures of Chloroperoxidase with Its Bound Substrates and Complexed with Formate, Acetate, and Nitrate. *J. Biol. Chem.* **2006**, *281*, 23990-23998. [CrossRef]

77. Sundaramoorthy, M.; Terner, J.; Poulos, T.L. Stereochemistry of the Chloroperoxidase Active Site: Crystallographic and Molecular-Modeling Studies. *Chem. Biol.* **1998**, *5*, 461-473. [CrossRef]

78. Shaik, S.; Kumar, D.; De Visser, S.P. A Valence Bond Modeling of Trends in Hydrogen Abstraction Barriers and Transition States of Hydroxylation Reactions Catalyzed by Cytochrome P450 Enzymes. *J. Am. Chem. Soc.* **2008**, *130*, 10128-10140. [CrossRef] [PubMed]

79. Hofrichter, M.; Ullrich, R. Heme-Thiolate Haloperoxidases: Versatile Biocatalysts with Biotechnological and Environmental Significance. *Appl. Microbiol. Biotechnol.* **2006**, *71*, 276-288. [CrossRef] [PubMed]

80. Farhangrazi, Z.S.; Sinclair, R.; Yamazaki, I.; Powers, L.S. Haloperoxidase Activity of Phanerochaete Chrysosporium Lignin Peroxidases H2 and H8. *Biochemistry* **1992**, *31*, 10763-10768. [CrossRef]

81. Ullrich, R.; Nueske, J.; Scheibner, K.; Spantzel, J.; Hofrichter, M. Novel Haloperoxidase from the Agaric Basidiomycete. *Appl. Environ. Microbiol.* **2004**, *70*, 4575-4581. [CrossRef]

82. Piontek, K.; Ullrich, R.; Liers, C.; Diederichs, K.; Plattner, D.A.; Hofrichter, M. Crystallization of a 45 KDa Peroxynase/Peroxidase from the Mushroom Agrocybe Aegerita and Structure Determination by SAD Utilizing Only the Haem Iron. *Acta Crystallogr. Sect. F: Struct. Biol. Cryst. Commun.* **2010**, *66*, 693-698. [CrossRef] [PubMed]

83. Zamocky, M.; Obinger, C. *Biocatalysis Based on Heme Peroxidases*; Torres, E., Ayala, M., Eds.; Springer: Berlin/Heidelberg, Germany, 2010; ISBN 978-3-642-12626-0.

84. China, H.; Okada, Y.; Dohi, T. The Multiple Reactions in the Monochlorodimedone Assay: Discovery of Unique Dehalolactonizations under Mild Conditions. *Asian J. Org. Chem.* **2015**, *4*, 1065-1074. [CrossRef]
85. Conesa, A.; Punt, P.J.; Van Den Hondel, C.A.M.J.J. Fungal Peroxidases: Molecular Aspects and Applications. *J. Biotechnol.* **2002**, *93*, 143–158. [CrossRef]

86. Cherry, J.R.; Lamsa, M.H.; Schneider, P.; Vind, J.; Svendsen, A.; Jones, A.; Pedersen, A.H. Directed Evolution of a Fungal Peroxidase. *Nat. Biotechnol.* **1999**, *17*, 379–384. [CrossRef]

87. Vilter, H. Peroxidases from Phaeophyceae: A Vanadium(V)-Dependent Peroxidase from Ascophyllum Nodosum. *Phytochemistry* **1984**, *23*, 1387–1390. [CrossRef]

88. Agarwal, V.; Miles, Z.D.; Winter, J.M.; Eustáquio, A.S.; el Gamal, A.A.; Moore, B.S. Enzymatic Halogenation and Dehalogenation Reactions: Pervasive and Mechanistically Diverse. *Chem. Rev.* **2017**, *117*, 5619–5674. [CrossRef]

89. Bertini, I.; Sigel, A.; Sigel, H. *Handbook of Metalloproteins*; Bertini, I., Sigel, A., Sigel, H., Eds.; John Wiley & Sons, Ltd.: New York, NY, USA, 2006; Volume 67, ISBN 0-8247-0520-3.

90. Leblanc, C.; Vilter, H.; Fournier, J.B.; Delage, L.; Potin, P.; Rebuffet, E.; Michel, G.; Solari, P.L.; Feiters, M.C.; Czjzek, M. Vanadium Haloperoxidases: From the Discovery 30 Years Ago to X-ray Crystallographic and V K-Edge Absorption Spectroscopic Studies. *Coord. Chem. Rev.* **2015**, *301–302*, 134–146. [CrossRef]

91. Messerschmidt, A.; Wever, R. X-ray Structure of a Vanadium-Containing Enzyme: Chloroperoxidase from the Fungus Curvularia Inaequalis. *Proc. Natl. Acad. Sci. USA* **1996**, *93*, 392–396. [CrossRef]

92. Fournier, J.-B.; Leblanc, C. Halogenation and Vanadium Haloperoxidases. In *Outstanding Marine Molecules*; La Barre, S., Kornprobst, J.-M., Eds.; Wiley-VCH Verlag GmbH & Co. KGaA: Weinheim, Germany, 2014; pp. 225–242. ISBN 9783527334650.

93. Neuwald, A.F. An Unexpected Structural Relationship between Integral Membrane Phosphatases and Soluble Haloperoxidases. *Protein Sci.* **1997**, *6*, 1764–1767. [CrossRef]

94. Renirie, R.; Hemrika, W.; Wever, R. Peroxidase and Phosphatase Activity of Active-Site Mutants of Vanadium Chloroperoxidase from the Fungus Curvularia Inaequalis. Implications for the Catalytic Mechanisms. *J. Biol. Chem.* **2000**, *275*, 11650–11657. [CrossRef]

95. Hasan, Z.; Renirie, R.; Kerkman, R.; Ruijssenaars, H.J.; Hartog, A.F.; Wever, R. Laboratory-Evolved Vanadium Chloroperoxidase Exhibits 100-Fold Higher Halogenating Activity at Alkaline PH: Catalytic Effects from First and Second Coordination Sphere Mutations. *J. Biol. Chem.* **2006**, *281*, 9738–9744. [CrossRef]

96. Fournier, J.B.; Rebuffet, E.; Delage, L.; Grijol, R.; Meslet-Cladière, L.; Rzonca, J.; Potin, P.; Michel, G.; Czjzek, M.; Leblanc, C. The Vanadium Iodoperoxidase from the Marine Flavobacteriaceae Species Zobellia Galactanivorans Reveals Novel Molecular and Evolutionary Features of Halide Specificity in the Vanadium Haloperoxidase Enzyme Family. *Appl. Environ. Microbiol.* **2014**, *80*, 7561–7573. [CrossRef]

97. Butler, A.; Carter-Franklin, J.N. The Role of Vanadium Bromoperoxidase in the Biosynthesis of Halogenated Marine Natural Products. *Nat. Prod. Rep.* **2004**, *21*, 180–188. [CrossRef]

98. Carter-Franklin, J.N.; Butler, A. Vanadium Bromoperoxidase-Catalyzed Biosynthesis of Halogenated Marine Natural Products. *J. Am. Chem. Soc.* **2004**, *126*, 15060–15066. [CrossRef]

99. Bar-Nun, N.; Shcolnick, S.; Mayer, A.M. Presence of a Vanadium-Dependent Haloperoxidase in Botrytis Cinerea. *FEMS Microbiol. Lett.* **2002**, *217*, 121–124. [CrossRef]

100. Plat, H.; Krenn, B.E.; Wever, R. The Bromoperoxidase from the Lichen Xanthoria Parietina Is a Novel Vanadium Enzyme. *Biochem. J.* **1987**, *248*, 277–279. [CrossRef]

101. Chen, Z. Recent Development of Biomimetic Halogenation Inspired by Vanadium Dependent Haloperoxidase. *Coord. Chem. Rev.* **2022**, *457*, 214404. [CrossRef]

102. Dong, J.J.; Ferná Ndez-Fueyo, E.; Li, J.; Guo, Z.; Renirie, R.; Wever, R.; Hollmann, F. Halofunctionalization of Alkenes by Vanadium Chloroperoxidase from Curvularia Inaequalis. *Chem. Commun.* **2017**, *53*, 6207. [CrossRef]

103. Naapuri, J.M.; Wagner, P.K.; Hollmann, F.; Deska, J. Enzymatic Bromocyclization of A- and Γ-Allenols by Chloroperoxidase from *Curvularia inaequalis*. *ChemistryOpen* **2022**, *11*, e202100236. [CrossRef]

104. Fernández-Fueyo, E.; van Wingerden, M.; Renirie, R.; Wever, R.; Ni, Y.; Holtmann, D.; Hollmann, F. Chemoenzymatic Halogenation of Phenols by Using the Haloperoxidase from Curvularia Inaequalis. *ChemCatChem* **2015**, *7*, 4035–4038. [CrossRef]

105. Wiesner, W.; van Pée, K.H.; Lingens, F. Purification and Characterization of a Novel Bacterial Non-Heme Chloroperoxidase from Pseudomonas Pyrrocinia. *J. Biol. Chem.* **1988**, *263*, 13725–13732. [CrossRef]

106. Hofmann, B.; Tölzer, S.; Pelletier, I.; Altenbuchner, J.; van Pée, K.H.; Hecht, H.J. Structural Investigation of the Cofactor-Free Chloroperoxidases. *J. Mol. Biol.* **1998**, *279*, 889–900. [CrossRef]

107. Pelletier, I.; Altenbuchner, J.; Mattes, R. A Catalytic Triad Is Required by the Non-Heme Haloperoxidases to Perform Halogenation. *Biochim. Biophys. Acta (BBA)/Protein Struct. Mol. 1995*, *1250*, 149–157. [CrossRef]

108. Picard, M.; Gross, J.; Lübbert, E.; Tölzer, S.; Krauss, S.; Van Pée, K.H.; Berkessel, A. Metal-Free Bacterial Haloperoxidases as Unusual Hydrolases: Activation of H₂O₂ by the Formation of Peracetic Acid. *Angew. Chem. Int. Ed. Engl.* **1997**, *36*, 1196–1199. [CrossRef]

109. Xu, G.; Wang, B.G. Independent Evolution of Six Families of Halogenating Enzymes. *PLoS ONE* **2016**, *11*, 1–13. [CrossRef]

110. Hecht, H.J.; Sobek, H.; Haag, T.; Pfeifer, O.; van Pée, K.-H. The Metal-Ion-Free Oxidoreductase from Streptomyces Aureofaciens Has an α/β Hydrolase Fold. *Nat. Struct. Biol.* **1994**, *1*, 532–537. [CrossRef]

111. Dairi, T.; Nakano, T.; Aisaka, K.; Katsumata, R.; Hasegawa, M. Cloning and Nucleotide Sequence of the Gene Responsible for Chlorination of Tetracycline. *Biosci. Biotechnol. Biochem.* **1995**, *59*, 1099–1106. [CrossRef]
85. Conesa, A.; Punt, P.J.; Van Den Hondel, C.A.M.J.J. Fungal Peroxidases: Molecular Aspects and Applications. *J. Biotechnol.* **2002**, *93*, 143–158. [CrossRef]

86. Cherry, J.R.; Lamsa, M.H.; Schneider, P.; Vind, J.; Svendsen, A.; Jones, A.; Pedersen, A.H. Directed Evolution of a Fungal Peroxidase. *Nat. Biotechnol.* **1999**, *17*, 379–384. [CrossRef]

87. Vilter, H. Peroxidases from Phaeophyceae: A Vanadium(V)-Dependent Peroxidase from Ascophyllum Nodosum. *Phytochemistry* **1984**, *23*, 1387–1390. [CrossRef]

88. Agarwal, V.; Miles, Z.D.; Winter, J.M.; Eustáquio, A.S.; el Gamal, A.A.; Moore, B.S. Enzymatic Halogenation and Dehalogenation Reactions: Pervasive and Mechanistically Diverse. *Chem. Rev.* **2017**, *117*, 5619–5674. [CrossRef]

89. Bertini, I.; Sigel, A.; Sigel, H. *Handbook of Metalloproteins*; Bertini, I., Sigel, A., Sigel, H., Eds.; John Wiley & Sons, Ltd.: New York, NY, USA, 2006; Volume 67, ISBN 0-8247-0520-3.

90. Leblanc, C.; Vilter, H.; Fournier, J.B.; Delage, L.; Potin, P.; Rebuffet, E.; Michel, G.; Solari, P.L.; Feiters, M.C.; Czjzek, M. Vanadium Haloperoxidases: From the Discovery 30 Years Ago to X-ray Crystallographic and V K-Edge Absorption Spectroscopic Studies. *Coord. Chem. Rev.* **2015**, *301–302*, 134–146. [CrossRef]

91. Messerschmidt, A.; Wever, R. X-ray Structure of a Vanadium-Containing Enzyme: Chloroperoxidase from the Fungus Curvularia Inaequalis. *Proc. Natl. Acad. Sci. USA* **1996**, *93*, 392–396. [CrossRef]

92. Fournier, J.-B.; Leblanc, C. Halogenation and Vanadium Haloperoxidases. In *Outstanding Marine Molecules*; La Barre, S., Kornprobst, J.-M., Eds.; Wiley-VCH Verlag GmbH & Co. KGaA: Weinheim, Germany, 2014; pp. 225–242. ISBN 9783527334650.

93. Neuwald, A.F. An Unexpected Structural Relationship between Integral Membrane Phosphatases and Soluble Haloperoxidases. *Protein Sci.* **1997**, *6*, 1764–1767. [CrossRef]

94. Renirie, R.; Hemrika, W.; Wever, R. Peroxidase and Phosphatase Activity of Active-Site Mutants of Vanadium Chloroperoxidase from the Fungus Curvularia Inaequalis. Implications for the Catalytic Mechanisms. *J. Biol. Chem.* **2000**, *275*, 11650–11657. [CrossRef]

95. Hasan, Z.; Renirie, R.; Kerkman, R.; Ruijssenaars, H.J.; Hartog, A.F.; Wever, R. Laboratory-Evolved Vanadium Chloroperoxidase Exhibits 100-Fold Higher Halogenating Activity at Alkaline PH: Catalytic Effects from First and Second Coordination Sphere Mutations. *J. Biol. Chem.* **2006**, *281*, 9738–9744. [CrossRef]

96. Fournier, J.B.; Rebuffet, E.; Delage, L.; Grijol, R.; Meslet-Cladière, L.; Rzonca, J.; Potin, P.; Michel, G.; Czjzek, M.; Leblanc, C. The Vanadium Iodoperoxidase from the Marine Flavobacteriaceae Species Zobellia Galactanivorans Reveals Novel Molecular and Evolutionary Features of Halide Specificity in the Vanadium Haloperoxidase Enzyme Family. *Appl. Environ. Microbiol.* **2014**, *80*, 7561–7573. [CrossRef]

97. Butler, A.; Carter-Franklin, J.N. The Role of Vanadium Bromoperoxidase in the Biosynthesis of Halogenated Marine Natural Products. *Nat. Prod. Rep.* **2004**, *21*, 180–188. [CrossRef]

98. Carter-Franklin, J.N.; Butler, A. Vanadium Bromoperoxidase-Catalyzed Biosynthesis of Halogenated Marine Natural Products. *J. Am. Chem. Soc.* **2004**, *126*, 15060–15066. [CrossRef]

99. Bar-Nun, N.; Shcolnick, S.; Mayer, A.M. Presence of a Vanadium-Dependent Haloperoxidase in Botrytis Cinerea. *FEMS Microbiol. Lett.* **2002**, *217*, 121–124. [CrossRef]

100. Plat, H.; Krenn, B.E.; Wever, R. The Bromoperoxidase from the Lichen Xanthoria Parietina Is a Novel Vanadium Enzyme. *Biochem. J.* **1987**, *248*, 277–279. [CrossRef]

101. Chen, Z. Recent Development of Biomimetic Halogenation Inspired by Vanadium Dependent Haloperoxidase. *Coord. Chem. Rev.* **2022**, *457*, 214404. [CrossRef]

102. Dong, J.J.; Ferná Ndez-Fueyo, E.; Li, J.; Guo, Z.; Renirie, R.; Wever, R.; Hollmann, F. Halofunctionalization of Alkenes by Vanadium Chloroperoxidase from Curvularia Inaequalis. *Chem. Commun.* **2017**, *53*, 6207. [CrossRef]

103. Naapuri, J.M.; Wagner, P.K.; Hollmann, F.; Deska, J. Enzymatic Bromocyclization of A- and Γ-Allenols by Chloroperoxidase from *Curvularia inaequalis*. *ChemistryOpen* **2022**, *11*, e202100236. [CrossRef]

104. Fernández-Fueyo, E.; van Wingerden, M.; Renirie, R.; Wever, R.; Ni, Y.; Holtmann, D.; Hollmann, F. Chemoenzymatic Halogenation of Phenols by Using the Haloperoxidase from Curvularia Inaequalis. *ChemCatChem* **2015**, *7*, 4035–4038. [CrossRef]

105. Wiesner, W.; van Pée, K.H.; Lingens, F. Purification and Characterization of a Novel Bacterial Non-Heme Chloroperoxidase from Pseudomonas Pyrrocinia. *J. Biol. Chem.* **1988**, *263*, 13725–13732. [CrossRef]

106. Hofmann, B.; Tölzer, S.; Pelletier, I.; Altenbuchner, J.; van Pée, K.H.; Hecht, H.J. Structural Investigation of the Cofactor-Free Chloroperoxidases. *J. Mol. Biol.* **1998**, *279*, 889–900. [CrossRef]

107. Pelletier, I.; Altenbuchner, J.; Mattes, R. A Catalytic Triad Is Required by the Non-Heme Haloperoxidases to Perform Halogenation. *Biochim. Biophys. Acta (BBA)/Protein Struct. Mol. 1995*, **1250**, 149–157. [CrossRef]

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128. Gkotsi, D.S.; Dhaliwal, J.; McLachlan, M.M.; Mulholand, K.R.; Goss, R.J. Halogenases: Powerful Tools for Biocatalysis (Mechanisms Applications and Scope). *Curr. Opin. Chem. Biol.* **2018**, *43*, 119–126. [CrossRef]

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118. Andorfer, C.; Lewis, J. Understanding and Improving the Activity of Flavin Dependent Halogenases via Random and Targeted Mutagenesis. *Annu. Rev. Biochem.* **2018**, *87*, 159–185. [CrossRef]

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125. Shinohara, C.; Chikanishi, T.; Nakashima, S.; Hashimoto, A.; Hamanaka, A.; Endo, A.; Hasumi, K. Enhancement of Fibrinolytic Activity of Vascular Endothelial Cells by Chaetoglobosin A, Crinipellin B, Geodin and Triticone B. *J. Antibiot.* **2000**, *53*, 262–268. [CrossRef]

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127. Chankhamjon, P.; Tsunematsu, Y.; Ishida-Ito, M.; Sasa, Y.; Meyer, F.; Boettger-Schmidt, D.; Urbansky, B.; Menzel, K.D.; Scherlach, K.; Watanabe, K.; et al. Regioselective Dichlorination of a Non-Activated Aliphatic Carbon Atom and Phenolic Bismethylation by a Multifunctional Fungal Flavoenzyme. *Angew. Chem.-Int. Ed.* **2016**, *55*, 11955–11959. [CrossRef]

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129. Frese, M.; Sewald, N. Enzymatic Halogenation of Tryptophan on a Gram Scale. *Angew. Chem.-Int. Ed.* **2015**, *54*, 298–301. [CrossRef]

130. Büchler, J.; Papadopoulou, A.; Buller, R. Recent Advances in Flavin-Dependent Halogenase Biocatalysis: Sourcing, Engineering, and Application. *Catalysts* **2019**, *19*, 1030. [CrossRef]

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132. Payne, J.T.; Poor, C.B.; Lewis, J.C. Directed Evolution of Rebh for Site-Selective Halogenation of Large Biologically Active Molecules. *Angew. Chem.-Int. Ed.* **2015**, *54*, 4226–4230. [CrossRef]

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134. Glenn, W.S.; Nims, E.; O'Connor, S.E. Reengineering a Tryptophan Halogenase to Preferentially Chlorinate a Direct Alkaloid Precursor. *J. Am. Chem. Soc.* **2011**, *133*, 19346–19349. [CrossRef]

135. Fraley, A.E.; Sherman, D.H. Enzyme Evolution in Fungal Indole Alkaloid Biosynthesis. *FEBS J.* **2020**, *287*, 1381–1402. [CrossRef]

136. Hillwig, M.L.; Liu, X. A New Family of Iron-Dependent Halogenases Acts on Freestanding Substrates. *Nat. Chem. Biol.* **2014**, *10*, 921–923. [CrossRef]

137. Matthews, M.; Krest, C.; Barr, E.; Vaillancourt, F.; Walsh, C.; Green, M.; Krebs, C.; Bollinger, J. Substrate-Triggered Formation and Remarkable Stability of the CH-Cleaving Chloroferryl Intermediate in the Aliphatic Halogenase, SyrB2. *Biochemistry* **2009**, *48*, 4331–4343. [CrossRef]

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182. Ngokpol, S.; Suwakulsiri, W.; Sureram, S.; Lirdprapamongkol, K.; Aree, T.; Wiyakrutta, S.; Mahidol, C.; Ruchirawat, S.; Kittakoop, P. Drimane Sesquiterpene-Conjugated Amino Acids from a Marine Isolate of the Fungus *Talaromyces minioluteus* (*Penicillium minioluteum*). *Mar. Drugs* **2015**, *13*, 3567–3580. [CrossRef] [PubMed]

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166. Amagata, T.; Tanaka, M.; Yamada, T.; Minoura, K.; Numata, A. Gymnastatins and Dankastatins, Growth Inhibitory Metabolites of a Gymnascella Species from a Halichondria Sponge. *J. Nat. Prod.* **2008**, *71*, 340–345. [CrossRef]

167. Konaklieva, M.I.; Dahl, M.L.; Turos, E. Halogenation Reactions of Epoxides. *Tetrahedron Lett.* **1992**, *33*, 7093–7096. [CrossRef]

168. Engler, M.; Anke, T.; Sterner, O.; Brandt, U. Pterulinic Acid and Pterulone, Two Novel Inhibitors of NADH: Ubiquinone Oxidoreductase (Complex I) Produced by a Pterula Species. I. Production, Isolation and Biological Activities. *J. Antibiot.* **1997**, *50*, 325–329. [CrossRef]

169. Kornsakulkarn, J.; Thongpanchang, C.; Chainoy, R.; Choowong, W.; Nithithanasilp, S.; Thongpanchang, T. Bioactive Metabolites from Cultures of Basidiomycete Favolaschia Tonkinensis. *J. Nat. Prod.* **2010**, *73*, 759–762. [CrossRef]

170. Lemaire, P.; Balme, G.; Desbordes, P.; Vors, J.P. Efficient Syntheses of Pterulone, Pterulone B and Related Analogues. *Org. Biomol. Chem.* **2003**, *1*, 4209–4219. [CrossRef]

171. Almeida, C.; Pérez-Victoria, I.; González-Menéndez, V.; de Pedro, N.; Martín, J.; Crespo, G.; Mackenzie, T.; Cautain, B.; Reyes, F.; Vicente, F.; et al. Non-Geminal Aliphatic Dihalogenation Pattern in Dichlorinated Diaporthins from Hamigera Fusca NRRL 35721. *J. Nat. Prod.* **2018**, *81*, 1488–1492. [CrossRef]

172. Cai, R.; Wu, Y.; Chen, S.; Cui, H.; Liu, Z.; Li, C.; She, Z. Peniisocoumarins A–J: Isocoumarins from Penicillium Commune QQF-3, an Endophytic Fungus of the Mangrove Plant Kandelia Candel. *J. Nat. Prod.* **2018**, *81*, 1376–1383. [CrossRef] [PubMed]

173. Bonini, C.; Righi, G. Regio- and Chemoselective Synthesis of Halohydrins by Cleavage of Oxiranes with Metal Halides. *Synthesis* **1994**, *1994*, 225–238. [CrossRef]

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175. Schafhauser, T.; Kirchner, N.; Kulik, A.; Huijbers, M.M.E.; Flor, L.; Caradec, T.; Fewer, D.P.; Gross, H.; Jacques, P.; Jahn, L.; et al. The Cyclochlorotine Mycotoxin Is Produced by the Nonribosomal Peptide Synthetase CctN in Talaromyces Islandicus ('Penicillium Islandicum'). *Environ. Microbiol.* **2016**, *18*, 3728–3741. [CrossRef]

176. Jiang, Y.; Ozaki, T.; Liu, C.; Igarashi, Y.; Ye, Y.; Tang, S.; Ye, T.; Maruyama, J.; Minami, A.; Oikawa, H. Biosynthesis of Cyclochlorotine: Identification of the Genes Involved in Oxidative Transformations and Intramolecular O,N-Transacylation. *Org. Lett.* **2021**, *23*, 2616–2620. [CrossRef]

177. Kropp, P.J.; Daus, K.A.; Crawford, S.D.; Tubergen, M.W.; Kepler, K.D.; Craig, S.L.; Wilson, V.P. Surface-Mediated Reactions. 1. Hydrohalogenation of Alkenes and Alkynes. *J. Am. Chem. Soc.* **1990**, *112*, 7433–7434. [CrossRef]

178. Wang, W.-X.; Cheng, G.-G.; Li, Z.-H.; Ai, H.-L.; He, J.; Li, J.; Feng, T.; Liu, J.-K. Curtachalasins, Immunosuppressive Agents from the Endophytic Fungus Xylaria Cf. Curta. *Org. Biomol. Chem.* **2019**, *17*, 7985–7994. [CrossRef]

179. Wang, W.-X.; Lei, X.; Yang, Y.-L.; Li, Z.-H.; Ai, H.-L.; Li, J.; Feng, T.; Liu, J.-K. Xylarichalasin A, a Halogenated Hexacyclic Cytochalasan from the Fungus Xylaria Cf. Curta. *Org. Lett.* **2019**, *21*, 6957–6960. [CrossRef]

180. Hemberger, Y.; Xu, J.; Wray, V.; Proksch, P.; Wu, J.; Bringmann, G. Pestalotiopens A and B: Stereochemically Challenging Flexible Sesquiterpene-Cycloaldic Acid Hybrids from *Pestalotiopsis* Sp. *Chem.-A Eur. J.* **2013**, *19*, 15556–15564. [CrossRef]

181. Wolpert, T.J.; Macko, V.; Acklin, W.; Jaun, B.; Arigoni, D. Structure of Minor Host-Selective Toxins From *Cochliobolus victoriae*. *Experientia* **1986**, *42*, 1296–1299. [CrossRef]

182. Ngokpol, S.; Suwakulsiri, W.; Sureram, S.; Lirdprapamongkol, K.; Aree, T.; Wiyakrutta, S.; Mahidol, C.; Ruchirawat, S.; Kittakoop, P. Drimane Sesquiterpene-Conjugated Amino Acids from a Marine Isolate of the Fungus *Talaromyces minioluteus* (*Penicillium minioluteum*). *Mar. Drugs* **2015**, *13*, 3567–3580. [CrossRef] [PubMed]

183. Kessler, S.C.; Zhang, X.; McDonald, M.C.; Gilchrist, C.L.M.; Lin, Z.; Rightmyer, A.; Solomon, P.S.; Turgeon, B.G.; Chooi, Y.-H. Victorin, the Host-Selective Cyclic Peptide Toxin from the Oat Pathogen Cochliobolus Victoriae, Is Ribosomally Encoded. *Proc. Natl. Acad. Sci. USA* **2020**, *117*, 24243–24250. [CrossRef] [PubMed]

184. Flatt, P.M.; O'Connell, S.J.; McPhail, K.L.; Zeller, G.; Willis, C.L.; Sherman, D.H.; Gerwick, W.H. Characterization of the Initial Enzymatic Steps of Barbamide Biosynthesis. *J. Nat. Prod.* **2006**, *69*, 938–944. [CrossRef] [PubMed]