This is the accepted version of the journal article: De, Sonakshi; Mattanovich, Diethard; Ferrer, Pau; [et al.]. «Established tools and emerging trends for the production of recombinant proteins and metabolites in Pichia pastoris». Essays in Biochemistry, Vol. 65, Issue 2 (July 2021), p. 293-307. DOI 10.1042/EBC20200138 This version is available at https://ddd.uab.cat/record/266632 under the terms of the license 1 Established tools and emerging trends for the production of recombinant 2 proteins and metabolites in Pichia pastoris 3 4 Sonakshi De1,2, Diethard Mattanovich1,2, Pau Ferrer3, Brigitte Gasser1,2,4,* 5 1 6 Institute of Microbiology and Microbial Biotechnology, Department of Biotechnology, 7 BOKU University of Natural Resources and Life Sciences Vienna, Austria 2 8 Austrian Centre of Biotechnology (ACIB), Vienna, Austria 3 9 Department of Chemical, Biological and Environmental Engineering, Universitat Autònoma 10 de Barcelona, Bellaterra (Cerdanyola del Vallès), Spain 4 11 Christian Doppler Laboratory for growth-decoupled protein production in yeast, BOKU 12 University of Natural Resources and Life Sciences Vienna, Austria 13 14 15 * Corresponding author: 16 Brigitte Gasser, Institute of Microbiology and Microbial Biotechnology, Department of 17 Biotechnology, BOKU University of Natural Resources and Life Sciences Vienna 18 Muthgasse 18, 1190 Vienna, Austria 19 brigitte.gasser@boku.ac.at 20 21 1 22 Abstract 23 Besides bakers’ yeast, the methylotrophic yeast Komagataella phaffii (also known as Pichia 24 pastoris) has been developed into the most popular yeast cell factory for the production of 25 heterologous proteins. Strong promoters, stable genetic constructs and a growing collection 26 of freely available strains, tools and protocols have boosted this development equally as 27 thorough genetic and cell biological characterization. This review provides an overview of 28 state-of-the-art tools and techniques for working with P. pastoris, as well as guidelines for 29 the production of recombinant proteins with a focus on small scale production for 30 biochemical studies and protein characterization. The growing applications of P. pastoris for 31 in vivo biotransformation and metabolic pathway engineering for the production of bulk and 32 specialty chemicals are highlighted as well. 33 2 34 Introduction 35 Methylotrophic yeasts raised the interest of biotechnologists first for their ability to grow on 36 methanol as the only carbon and energy source, thus promising the production of cheap 37 protein rich biomass for animal and human nutrition (1). The oil crisis in the 1970s 38 interrupted these plans, however, triggered the development of these yeasts as platforms 39 for the production of heterologous proteins. Among them, Komagataella phaffii (formerly 40 known as Pichia pastoris) stands out by its wide application in research labs and industrial 41 production. More recently, P. pastoris was also employed for the production of metabolites 42 by metabolic engineering. A well-annotated genome sequence (2) as well as genome editing 43 tools and collections of synthetic biology parts and devices make P. pastoris a promising 44 chassis for synthetic biology applications (3). 45 The first yeast strain of what is known today as the Komagataella genus was isolated in 1920 46 by Alexandre Guillermond from a wounded horse chestnut tree and described as 47 Zygosaccharomyces pastori (4). Further strains were described as Pichia pastoris by Herman 48 Phaff, isolated mainly from oak and pine trees of Southwest United States (5). Based on 49 ribosomal gene sequences P. pastoris was further allocated to a new genus, Komagataella, 50 and split into two species. With new isolates found over the following years Komagataella 51 comprises seven species today (6). The strains used in biotechnology however are usually 52 still all referred to as Pichia pastoris which will be used as a synonym for Komagataella spp. 53 in the following as well. 54 55 Methylotrophy 56 Methylotrophy, i.e. the ability to use methanol and similar one-carbon molecules as carbon 57 and energy source, has developed several times among bacteria and yeasts. Methylotrophic 58 yeasts are phylogenetically related and comprise species of the genera Komagataella, 59 Ogataea, and some Candida sp. (7). They share the specific metabolism, first oxidizing 60 methanol by alcohol oxidases (AOX) to formaldehyde, which is further assimilated by 61 dihydroxyacetone synthase (DAS) via the xylulose monophosphate cycle (Figure 1). Due to 62 the low specific activity of these enzymes extraordinarily high expression levels are required, 63 so that the promoters regulating AOX1 and the two DAS genes are stronger than any other 64 metabolic gene promoters of K. phaffii. The entire xylulose monophosphate cycle is localized 65 in peroxisomes which probably increases its efficiency by metabolic channeling and shielding 3 66 the cytosol from toxic intermediates like hydrogen peroxide (8, 9). In three interconnected 67 cyclic pathways 3 xylulose-5-phosphate molecules are regenerated while one 68 glyceraldehyde-3-phosphate is built as a precursor for biomass formation (Figure 1). This 69 pathway shares surprising similarity to the Calvin-Benson-Bessham cycle of CO2 assimilation 70 (9). Besides this assimilatory pathway, energy is produced by oxidation of formaldehyde to 71 CO2 via formate, yielding cytosolic NADH which is both used as reduction equivalent and 72 channeled into the respiratory chain to provide ATP. 73 Methylotrophy was first employed in biotechnology for the production of yeast biomass 74 from natural gas for food and feed applications (1), but soon also considered useful for 75 recombinant protein production because of the strong and regulated promoters, mainly the 76 AOX1 promoter of P. pastoris (10). In a standard setup, methanol is used both as the carbon 77 and energy source and as the inducer of heterologous gene expression. To this end, three 78 different methanol utilization (Mut) phenotypes are employed based on the presence or 79 deletion of the two alcohol oxidase genes. The wild type with both AOX1 and AOX2 active is 80 called Mut+, while deletion of the main form, AOX1, leads to the MutS (methanol utilization 81 slow) phenotype, and strains deleted in both AOX genes are called Mut-. MutS strains are 82 often employed in protein production while Mut- strains are considered less suitable based 83 on the assumption that they lack any energy supply from methanol, and also do not provide 84 for formaldehyde and formate which are considered to play a major role in the induction of 85 methanol utilization genes (11). Zavec et al. (12) showed however that also Mut- strains 86 consume methanol and can serve as a platform for the production of proteins. The same 87 authors demonstrated recently that native alcohol dehydrogenase (Adh2) of P. pastoris is 88 responsible for methanol oxidation in Mut- strains (13). 89 90 Tools & Techniques 91 From strain construction until bioprocess optimization for recombinant production in any 92 expression system, several product-specific aspects need to be considered. This is where the 93 well-established and constantly expanding set of tools and techniques available for strain 94 and bioprocess engineering (Figure 2) contributes in making P. pastoris a highly efficient 95 expression system (14, 15). 96 97 4 98 Design of the expression construct 99 P. pastoris offers a range of well-characterized constitutive and inducible promoters to 100 choose from. The constitutive promoter PGAP has frequently been used in large scale 101 production of recombinant proteins in P. pastoris (16). Other strong constitutive promoters 102 include PTEF1 and PGCW14 (17). However, constitutive promoters are not ideal for recombinant 103 proteins that are toxic to the cells. In terms of inducible promoters, most commonly the 104 methylotrophic feature of P. pastoris is harnessed by driving the expression of recombinant 105 genes under the influence of the methanol-inducible promoter of the alcohol oxidase I 106 (AOX1) gene (18). Other strong methanol-based promoters include PFLD1 (also induced by 107 methylamine), PDAS and PCAT1 (also induced by glycerol and de-repressed under carbon 108 source limitation) (17). While methanol-based promoters provide strong induction, their 109 disadvantage in terms of toxicity of methanol has led to extensive research for the discovery 110 of alternative, methanol-independent expression systems. On one hand, cell engineering has 111 been successfully applied to develop methanol-independent PAOX1 and PDAS1 host systems 112 (19, 20). Hartner et al. created a library of several PAOX1 variants, by which they not only 113 identified variants with activities higher than the native promoter upon methanol induction, 114 but also identified some variants which demonstrated strong methanol-independent activity 115 in de-repressing conditions (21). On the other hand, several methanol-independent 116 promoters were identified, including PGTH1 (induced in glucose-limited conditions), PTHI11 117 (thiamine repressed) and PADH3 (ethanol-induced) (22-24). Orthologous promoters have also 118 been successfully used, for example, the promoter of the methanol oxidase gene from 119 Hansenula polymorpha (PMOX) (25, 26). Promoter engineering has been applied to some of 120 these native promoters as effective tool for engineering existing promoters for better 121 performance (27-30). Additionally, bidirectional promoters have been engineered which can 122 offer an advantage when co-expression of multiple genes is necessary (31). 123 124 Secretion of the recombinant product can simplify the downstream purification steps and 125 therefore is often desirable. Signal peptides play a very important role in efficiently driving 126 the secretion of the recombinant protein. While the secretion signal of the Saccharomyces 127 cerevisiae α-factor mating pheromone is most commonly used to drive the secretion of 128 recombinant proteins in P. pastoris, there have been instances where the native signal 129 peptide of the protein or other endogenous signal peptides were shown to have similar or 5 130 even higher efficiencies (32, 33). Additionally, higher yields of recombinant proteins have 131 been demonstrated using modified versions of the S. cerevisiae α-mating factor secretion 132 signal (34). However, selection of signal peptide can be protein-dependent and therefore, 133 testing multiple signal peptides can lead to higher yields (35). 134 In addition to transcriptional termination, terminator regions can also influence the level of 135 expression. The toolbox of terminators in P. pastoris was quite small, consisting of only 20 136 terminators with not much variation in expression between them (36). Only recently, a 137 catalog of 72 endogenous, heterogeneous and synthetic terminators was developed and 138 characterized, within which a tunability of 17-fold was observed (37). 139 140 Selection of vector and background strain 141 While there have been some studies towards the generation of circular plasmids with 142 autonomously replicating sequences (ARS) for stable recombinant expression in P. pastoris, 143 genomic integration vector systems are most commonly used (38, 39). A selection of vector 144 systems is commercially available with a choice of antibiotic or auxotrophic selection 145 markers (40, 41). Additionally, there are vectors based on the GoldenPiCS modular cloning 146 system which enable single reaction assembly of selected promoters, terminators, resistance 147 cassettes and genomic integration loci (36). For selection of a background strain, one gets to 148 choose from various wild-type and auxotrophic strains, strains with different methanol- 149 utilization phenotypes (Mut+, MutS & Mut-), protease-deficient strains and glycol-engineered 150 strains (42). 151 152 Molecular biology methods 153 Several well-established molecular biology techniques in P. pastoris aid in speeding up the 154 strain generation process (Figure 2). The modular cloning toolbox of GoldenPiCS, mentioned 155 earlier, allows one-step assembly of multiple expression cassettes into a single vector, which 156 can then be linearized and integrated into the genome of the host strain. This method 157 requires the use of selection markers, which can be exploited to increase the copy number 158 of the recombinant gene construct by increasing the selection pressure. This can be an 159 advantage since increased gene dosage is often associated with increase in protein titers (15, 160 43, 44). In case of antibiotic resistance markers, there is a possibility for removal of the 161 resistance gene post selection of positive clones by transient expression of Cre recombinase, 6 162 however, this is not applicable if multicopy strains are intended (45). Recent development 163 and fine-tuning of the CRISPR/Cas9 technology in P. pastoris allows for marker free 164 integration of expression cassettes (46-48). While the technology still has its limitations, a 165 large amount of research has been directed in improving the technology leading to higher 166 efficiencies, possibility of multiple genomic integrations as well as application of deactivated 167 Cas9 (dCas9) for targeted gene interference (49). 168 169 Strain selection & bioprocess optimization 170 The final steps in strain development comprise evaluation of generated strains for final clone 171 selection and bioprocess optimization (Figure 2). Strain evaluation, often also referred to as 172 “screening”, is usually done in shake-flasks or in high-throughput microtiter plate cultivation. 173 Guidelines for such screenings are given in Box 1. To find suitable production clones, 174 screening of at least 20-40 clones is recommended. Depending on the protein of interest and 175 the desired product titer screening of several hundred clones may be required, which makes 176 high-throughput screening methods desirable. While on one hand it is possible to mimic 177 batch or fed-batch fermentation conditions and select optimum clones, bioprocess 178 optimization is usually not possible in shake-flasks/microtiter plates. In that respect 179 mini/microscale cultivations or microfluidics and Lab-on-Chip based technologies provide 180 alternative screening platforms that also integrate bioprocess optimization, making scale-up 181 more convenient (50-53). 182 183 Strain engineering to improve protein production 184 Several studies report strategies on how to enhance recombinant protein production and 185 secretion in P. pastoris (summarized in 54, 55). Such cell engineering approaches are mostly 186 not applied for initial characterization, unless a specific bottleneck is observed. Examples are 187 mainly the avoidance of proteolytic degradation through use of strains deficient in the major 188 cellular proteases such as pep4 deficient strains or the co-expression of certain chaperones, 189 especially protein disulfide isomerase (55, 56). Some proteins such as cytochrome P450s or 190 heme-containing proteins require co-factors that either need to be added externally or 191 produced by the cells (57). 192 193 7 194 Systems biology 195 While a lot of advances has been made regarding the tools and techniques for strain 196 engineering in P. pastoris, development of systems biology tools is steadily gaining 197 importance. For example, genomic and proteomic studies have been applied to some 198 recombinant strains under conditions of recombinant protein production (54). Metabolic 199 flux analysis (MFA) has been carried out to understand the metabolic impact of recombinant 200 protein production in P. pastoris (58). Extensive MFA studies have been performed to 201 compare metabolite concentrations under different cultivation conditions (9, 59, 60). 202 203 Genome scale metabolic models (GEMs) have the potential to identify engineering targets to 204 alleviate metabolic burdens as well as to develop efficient bioprocess strategies. Several 205 GEMs have been available for P. pastoris since 2010 (61-65). Additionally, efforts have been 206 directed towards refining or upgrading existing GEMs, for e.g. by incorporating native and 207 humanized N-glycosylation pathways for production of glycoproteins (66), improving 208 biomass synthesis equations allowing improved prediction capabilities over a wide range of 209 substrates (67) or including additional reaction pathways (68). Thus, systems-level studies 210 are now being incorporated into the strain engineering and improvement workflow. 211 212 Pichia pastoris as a host for recombinant protein production 213 214 P. pastoris produced recombinant proteins for biomedical and industrial applications 215 Since the late 1990s P. pastoris has been used to produce proteins for biopharmaceutical, 216 industrial and diagnostic applications. Commercialized products include insulin, growth 217 factors, interferon, and subunit vaccines such as hepatitis B surface antigen, and several 218 others including peptides and antibody fragments (especially single chain antibody 219 fragments and nanobodies) are in the clinical pipeline (56, 69, 70). Furthermore, industrial 220 enzymes such as phospholipase C and phytase are produced using P. pastoris as host (71, 221 72). P. pastoris is also described to be a preferred expression system to prepare several 222 other recombinant subunit antigens against human and animal pathogens (70, 73), e.g. 223 brucellosis subunit vaccine (74), Zika virus envelope domain III (75), influenza hemagglutinin 224 and neuraminidase (69), and lately also Sars-CoV-2 spike antigen (76). 8 225 Very recently, P. pastoris has entered into another emerging market, as soy leghemoglobin 226 produced in P. pastoris was approved as a flavor component of cultured meat by the FDA 227 (77, 78). 228 While virus antigens are often produced as intracellular virus like particles (VLPs) (73) or self- 229 assembling nanoparticles (69), most proteins produced in P. pastoris are secreted (56, 70). 230 This allows for a reduced number of downstream processing steps during purification 231 compared to intracellular production (56). As several proteins secreted by P. pastoris have 232 received a GRAS notice by the FDA (77), it is even possible to directly use the culture 233 supernatant containing the secreted protein of interest in some cases such as in animal feed 234 applications (79). 235 236 Production of recombinant proteins for structural and biophysical analysis 237 Apart from being an important host for commercial protein production purposes, a high 238 number of published studies deal with using P. pastoris to provide a wide variety of 239 heterologous proteins and mutants thereof for biochemical characterization such as enzyme 240 activity or for structural and biophysical studies. Often these studies are the first steps for 241 later utilization of enzymes. For example, P. pastoris was used to determine 1.7-Å resolution 242 crystal structures and to identify the substrate specificities and the catalytically active sites 243 of several Aspergillus rutinosidases, which are potential catalysts for flavonoid compounds 244 with nutritional value (80, 81). Substrate specificity and crystal structures of several plant 245 and fungal carbohydrate active enzymes have been elucidated by expressing them in P. 246 pastoris. Heterologous expression of plant glycosyltransferases did not only result in the 247 production of mannan and glucomannan in P. pastoris, but additionally provided important 248 insights into the yet unresolved biosynthesis of these polysaccharides in plants (82, 83). Site- 249 directed mutagenesis was also carried out for proteins of biomedical interest, e.g. to study 250 immunogenic residues of the buckwheat allergen (84), to increase efficacy and stability of 251 ocriplasmin (85) or to strengthen the affinity of a scFv to its target and should be even more 252 facilitated by the high-throughput cloning tools available for P. pastoris now (see section 253 Tools & Techniques). 254 255 Membrane proteins (MP) are a particularly challenging class of proteins that have been 256 produced with a quite high rate of success in P. pastoris (86, 87). Several efforts have been 9 257 undertaken to improve the expression levels and purification of MPs. Common cell 258 engineering strategies and tools for MP production in yeast were summarized by Byrne, 259 2015 and Routledge et al. 2016 (86, 88). Recently, a novel procedure was devised in which 260 misfolding and aggregation of integral MPs is avoided by directly solubilizing them from 261 protoplasts instead of crude membrane preparations (89). Nanodisc reconstitution was 262 successfully applied to generate sufficient amounts of purified channelrhodopsins for 263 biophysical characterization (90). The human tetraspanin CD81 expressed in P. pastoris could 264 be efficiently solubilized and purified within a lipid environment by using styrene-maleic 265 anhydride co-polymers, providing a platform to study the influence of protein-lipid 266 interactions of tetraspanins (91). 267 Crystal structures and biophysical characterization of human amino acid transporters, 268 aquaporins and several human G protein coupled receptors (GPCRs) expressed in P. pastoris 269 contributed significantly to our understanding of their substrate transport dynamics (87, 92- 270 94). The recent elucidation of the mating pathway in P. pastoris together with its high 271 expression capacity of GPCRs, so far mainly exploited for structural studies, paves the way to 272 perform in vivo GPCR signaling studies as described for S. cerevisiae (95, 96). 273 274 P. pastoris has been used since the 1990s to produce isotope-labeled proteins for NMR- 275 based structural analysis, and it is the preferred yeast host for this powerful structural 276 biology technique (87, 94). NMR spectroscopy is well suited to study protein-ligand 277 interactions and dynamics and to guide ligand design, however, for such purposes 278 incorporation of sophisticated isotopic labels is often required. Zhang, 2020 provided a 279 collection of suitable labeling protocols and summarized the most recent advances made in 13 15 19 280 P. pastoris (94). Labeling strategies include global or selective C N-labeling, F-labeling, 281 deuterium labeling for improved resolution of spectra, and methyl-labeling of valine, leucine 282 or isoleucine for even more improved sensitivity of spectra of high-molecular-weight 283 proteins or complexes. Site-specific methyl-labeling in a deuterated background permitted 284 sensitive methyl-TROSY experiments in P. pastoris that were successfully conducted to 285 elucidate structure-function relationships of several human GPCRs of biomedical relevance 286 (87). On the other hand, selenomethionine labeling might be applied to improve X-ray 287 crystallography (97). For all of the latter approaches, uptake of labeled amino acid(s) might 288 be limiting, thus cell engineering provides a promising strategy for improving labeling 10 289 efficiencies (94, 98). Cell-free expression systems recently established also for P. pastoris are 290 another alternative especially for the incorporation of otherwise cytotoxic selenomethionine 291 (99). 292 293 Design of culture conditions for small scale protein production 294 While projects aiming at producing a certain protein or compound usually required large 295 scale cultivation in bioreactors, studies aiming at protein characterization are often (at least 296 initially) performed in small scale such as shake flasks or deep well plates (100). Guidelines 297 for selecting suitable producer clones are given in Box 1. 298 Often, substantial efforts are invested to improve the cultivation conditions for a certain 299 protein of interest. Based on our experience we suggest investigating the impact of pH, 300 which should not be too close to the isoelectric point of the protein to avoid precipitation, 301 and may as well impact product quality and stability through proteolysis. Many researchers 302 investigate methanol concentration, with the outcome that in most cases addition of 0.5-1% 303 methanol twice a day works best. Usually higher cell density and higher product titers are 304 obtained in complex medium containing yeast extract and peptone or casamino acids, 305 however, proteolytic activity (probably due to cell lysis) might be observed. For secretory 306 proteins, growth-limiting carbon supply (e.g. by enzyme-mediated glucose release simulating 307 a fed batch regime) leads to good product titers and high product quality in our experience 308 (49, 101), and works comparably well for both complex and synthetic media. This cultivation 309 strategy is also well suited for the above described amino acid labeling strategies where the 310 use of minimal medium, with or without deuteration, is required, and helps to overcome the 311 previously reported limit of low protein amounts. 312 313 Metabolite production by whole-cell biotransformation and fermentation 314 bioprocesses 315 316 Although P. pastoris has been developed primarily as a cell factory for recombinant protein 317 production, its potential for metabolite production purposes has received increasing 318 attention over the past 15 years, likely boosted by the increasing physiological knowledge 319 base and synthetic biology tools availability (see excellent reviews by (102-104)). The use of 320 P. pastoris has been demonstrated for the production of diverse chemical compounds, 11 321 particularly for complex metabolites such as plant secondary metabolites (polyketides, 322 terpenoids, isoflavonoids) and other drug metabolites (e.g. steroids, sphingolipids) as well 323 as, to a much lesser extent, for bulk chemicals such as organic acids, biofuels or biopolymers 324 (Figure 3). Nonetheless, most of these examples are proof-of-principle studies reporting 325 titers, yields and productivities far from industrially attractive metrics. 326 Two major metabolite production approaches have been explored, namely 327 biotransformation (biocatalysis) and fermentation bioprocesses. Whereas in fermentations 328 the products are synthesized from carbon/nitrogen substrates via the host cells’ native or 329 engineered metabolism, in biotransformations, cell growth and product synthesis phase are 330 separated, i.e. biotransformations are typically performed by resting cells, which convert 331 substrates/precursors to the desired products. Generally, biotransformations are catalyzed 332 by intracellular enzymes, although cell surface display of enzymes for biotransformation 333 purposes has also been demonstrated. 334 P. pastoris presents several a priori physiological advantages and specific metabolic traits 335 that make this yeast particularly interesting for whole-cell biocatalytic systems. i) As 336 Crabtree negative yeast, high cell densities, that is, high amounts of biocatalyst, can be easily 337 obtained from bioreactor cultures using cheap substrates; also, P. pastoris shows higher 338 resistance to harsh process conditions such as low pH compared to other cell factories such 339 as Escherichia coli. ii) The native alcohol oxidase (Aox), a peroxisomal enzyme with relative 340 low substrate specificity, can catalyze the oxidation of many short-chain alcohols to the 341 respective aldehydes using oxygen as electron acceptor, generating hydrogen peroxide as a 342 toxic by-product. Notably, P. pastoris also synthesizes large amounts of catalase in the 343 peroxisomes together with Aox during methylotrophic growth, thereby providing an 344 interesting system for oxidase-catalase-based reactions (104). This concept can be further 345 expanded by, e.g. targeting heterologous oxidases such as D-amino acid oxidases (DAO) to 346 the peroxisome, providing a system for α-keto acids production from α-amino acid 347 substrates and resolution of racemic mixtures of amino acids (105). iii) The native methanol 348 dissimilation pathway, which oxidizes formaldehyde stepwise to formate and CO2 by two 349 NAD+-dependent dehydrogenases has been exploited as a natural NADH regeneration 350 system for reduction reactions. Moreover, such endogenous system has been specifically 351 optimized for whole-cell biotransformation purposes by overexpressing the formaldehyde 352 dehydrogenase encoding gene (FLD) and disrupting the DAS genes, key genes in the 12 353 competing methanol assimilatory pathway (106, 107). Also, NADPH regeneration engineered 354 systems or process strategies have been proposed (104). For instance, Tang and co-workers 355 (108) have used glucose as a co-substrate for NADPH synthesis to improve NADPH- 356 dependent synthesis of the steroid boldenone, an androgenic–anabolic steroid and a 357 testosterone derivative. iv) P. pastoris has shown a high aptitude for expression of 358 membrane bound proteins like cytochrome P450 monooxygenases (CYP) and reductases 359 (CPR). v) Under methanol-growing conditions, peroxisomes largely proliferate which offers 360 the possibility to compartmentalize synthetic metabolic pathways in this organelle as a 361 metabolic engineering strategy (103). 362 P. pastoris has also been used in whole-cell biotransformations requiring ATP such as the 363 synthesis of dipeptides and tripeptides, as well as in other reactions not dependent of 364 cofactors, e.g. hydrolysis, carbon-carbon bond formation reactions, and further synthesis 365 reactions. Some recent exemplary studies include the production of the dipeptides (109), 366 synthesis of phospholipids (110), and the use of transketolase-overproducing strains to 367 catalyze asymmetric carbon-carbon bond formation reactions, e.g. the production of L- 368 erythrulose from prochiral substrates (111) or, strains combining the overexpression of 369 transketolase and ω-transaminases encoding genes for chiral amino-alcohols synthesis (112, 370 113). 371 372 Despite the fact P. pastoris presents several features such as an industrially well-established 373 fermentation technology and the ability to grow on renewable feed stocks like glycerol and 374 methanol that are more reduced than glucose, it remains largely unexploited as a cell factory 375 for bulk/platform chemicals and biofuels production (102, 103). For instance, conversion of 376 glycerol into L-lactic acid has been demonstrated by the expression of a lactate 377 dehydrogenase gene from Bos taurus (114). Additional metabolic engineering of this strain 378 investigated the impact of deletion of the pyruvate decarboxylase gene, aiming at reducing 379 acetic acid formation and consequently obtaining higher lactic acid titers. This resulted in a 380 yield of 67% L-lactic acid and 20% arabitol as a by-product in glycerol batches with oxygen 381 limitation (115). Further disruption of the arabitol dehydrogenase encoding gene has been 382 recently reported by the same group (116), obtaining an increase of 20% in lactic acid and a 383 50% reduction in arabitol yields in chemostat cultures. Also recently, the production of D- 384 lactic acid from methanol using an engineered P. pastoris strain expressing multiple copies of 13 385 the D-lactate dehydrogenase gene from Leuconostoc mesenteroides has been reported 386 (117). Similarly, production of an increasing number of platform chemicals has also been 387 demonstrated, typically at shake flask or small bioreactor scale, including isobutanol and 388 isobutyl acetate (118), isopentanol (3-Methyl-1-butanol) (119), and 2,3-butanediol (120). 389 Moreover, the potential of P. pastoris to produce malic acid from methanol has been lately 390 explored through different metabolic engineering strategies, obtaining up to about 2.8 g/L of 391 malic acid in shake flask cultures (121). 392 393 So far, these novel developments to metabolically engineer P. pastoris for metabolite 394 production have generally relied on classical genetic tools (promoters, markers etc.) and 395 conventional strains that were originally optimized for recombinant protein production. 396 Despite enormous advances in systems and synthetic biology tools in recent years, 397 significant challenges in genome engineering remain compared to Saccharomyces cerevisiae, 398 e.g. generally lower efficiency of currently available CRISPR and homologous recombination 399 based methods (122). The incorporation of new-generation synthetic biology tools should 400 speed up development of metabolite production strains beyond mere demonstration/proof- 401 of-concept. To this end, the design of robust chassis strains supporting efficient conversion 402 of alternative carbon substrates/energy sources, for e.g., CO2 (123), acetate (124) or 403 cellulose/cellobiose (125) to key metabolic precursors e.g. acetyl-CoA, farnesyl 404 pyrophosphate, malonyl-CoA (Figure 3) will be key in the forthcoming years. 405 406 Summary points 407 • Methylotrophy and the strong methanol regulated promoters were the key drivers to 408 develop P. pastoris to a protein production host. 409 • Early free access to strains and vectors has promoted research on P. pastoris 410 enormously. 411 • The development of tools and parts for P. pastoris make it an important synthetic 412 biology chassis. 413 • A wide range of available molecular and synthetic biology toolkits contributes in 414 developing a simple workflow from strain design to bioprocess optimization. 415 • The efficient native one-carbon metabolism offers promising perspectives for a 416 single-carbon (methanol, formate, CO2) based bioeconomy. 14 417 418 Author contributions: All authors contributed to the manuscript and figure drafts, SD 419 prepared the final figures. All authors have read and agreed to the final version of the 420 manuscript. 421 Declaration of Interests: The authors declare no competing interests. 422 Funding: BG, SD and DM acknowledge funding by the Austrian BMDW and Nationalstiftung 423 FTE through the Christian Doppler Research Association, by BMK, BMDW, SFG, 424 Standortagentur Tirol, Government of Lower Austria und Vienna Business Agency through 425 the FFG – COMET Funding Program, and by the Austrian Science Fund FWF (BioToP, grant 426 W1224). PF acknowledges support from the Spanish Ministry of Science and Innovation: 427 Project PID2019-104666GB-I00, and from the Agency for Management of University and 428 Research Grants (AGAUR) of the Catalan Government: Grant 2017-SGR-1462 - Program to 429 support activities of research groups. 430 431 15 432 Figure Legends 433 Figure 1: Metabolic pathways of methanol utilization in P. pastoris. After oxidation of 434 methanol to formaldehyde assimilation is achieved in the xylulose-5-phosphate cycle, 435 indicated in green. A byproduct of methanol oxidation is hydrogen peroxide, which is further 436 detoxified by catalase. These reactions, as well as the Xu5P cycle, are localized in 437 peroxisomes. Formaldehyde is also dissimilated to CO2 in three cytosolic reactions, indicated 438 in blue. The produced NADH serves as reduction equivalents and for ATP production. 439 Assimilation of three formaldehyde molecules leads to the release of one molecule of 440 glyceraldehyde-3-phosphate from peroxisomes which serves for biomass growth. 441 GSH: reduced glutathione; GS-CH2OH: S-(hydroxymethyl)glutathione; GS-CHO: S- 442 formylglutathione; Cat1: catalase; Fld1: formaldehyde dehydrogenase; Fgh1: S- 443 formylglutathione hydrolase; Fdh1: formate dehydrogenase; Aox1/2: alcohol oxidase 1 and 444 2; Das1/2: dihydroxyacetone synthase 1 and 2; Dak2: dihydroxyacetone kinase; Tpi1: 445 triosephosphate isomerase; Fba1-2: fructose 1,6-bisphosphate aldolase; Fbp1: fructose 1,6- 446 bisphosphatase; Shb17: sedoheptulose 1,7-bisphosphatase; Rki1-2: Ribose 5-phosphate 447 ketol-isomerase; Rpe1-2: D-ribulose 5-phosphate 3-epimerase. 448 449 450 16 451 Figure 2: Workflow from construct design to bioprocess optimization and strain 452 improvement for recombinant protein production in P. pastoris. A. After selection of the 453 different modules, the expression cassette can be constructed using GoldenPiCS. There is a 454 possibility to integrate the expression construct in the recombinant strain either with 455 selection marker by direct plasmid linearization and integration or marker-free by 456 CRISPR/Cas9-based homology directed recombination. B. An optimum strain background 457 depending on the recombinant protein can be selected. C. Well-established transformation 458 protocols are available for generating the recombinant strain. D. Different screening 459 strategies enable evaluation of a large number of clones resulting in faster clone selection. 460 Lab-on-Chip technology also allows bioprocess optimization at this stage. E. Final clones can 461 be cultivated in bioreactors for producing high titers of the recombinant protein. F. Data 462 obtained during bioreactor cultivations combined with systems biology tools can be used to 463 identify further cell engineering targets for strain improvement 464 465 466 17 467 Figure 3: Metabolites overproduced in P. pastoris. Major pathways towards key metabolic 468 precursors or metabolic nodes (in brown) and products already made by P. pastoris (green) 469 are depicted in black. Heterologous pathways and metabolites are depicted in blue. Relevant 470 substrates that P. pastoris can naturally assimilate as carbon and/or energy source are 471 shown in yellow. 472 473 474 18 475 Box1: Recommendations for selection of a producer strain 476 Guidelines how to select strains for biochemical studies or protein production purposes. 477 1. After transformation, plate aliquots on a suitable selection medium e.g. YPD containing 478 100 µg/mL Zeocin (in case your vector has the Zeo resistance cassette). As strains with 479 higher gene copy number are often correlated with higher product titers, you might 480 also plate aliquots at a higher Zeocin concentration e.g. 250 µg/mL, 500 µg/mL or 1000 481 µg/mL. 482 2. After restreaking, use 20-40 transformants (selected from all Zeocin concentrations) 483 for the initial screening. On average, 5-10% of the clones outperform the 484 others. There may be cases where it is necessary to screen more clones, e.g. when 485 searching for a high production clone of a very difficult to produce protein. 486 3. Initial screening can be performed in 24 deep well plates, 96 deep well plates or any 487 other suitable format depending on the volume needed for quantification and analysis. 488 4. Select the pH of the cultivation medium according to your protein of interest (avoid 489 being close to the isoelectric point or a pH where degradation might occur). In most 490 cases, the starting pH of the medium is between 5.0-7.0, and a suitable buffer (e.g. 491 phosphate buffer, MES (2-(N-morpholino)ethanesulfonic acid), citrate) is used to 492 stabilize the pH during cell growth. 493 5. Prepare cryo stocks of your strains (e.g. 1 mL of selective overnight culture + 10% 494 glycerol and freeze at -70°C) early during screening. For further experiments, the 495 preparation of working cell banks (WCBs) from this stock is recommended to avoid 496 repeated freeze/thaw cycles of the stock. 497 6. Rescreen the 3-5 best or most-interesting clones to evaluate their performance, and 498 select 1 or 2 for further experiments. 499 7. If optimization of screening conditions is required, we recommend to evaluate 500 different pH values (see above) and cultivation temperatures (e.g. 20-25°C compared 501 to 28-30°C). 502 8. 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