Breaking the cell wall for efficient DNA delivery to diatoms

Domain: neuroscience
Published: 2026-01-22
Editorial outcome: minor_revision

Abstract
Diatoms produce 20% of the world’s fixed organic carbon yet remain under-utilized as cell factories due to limited genetic engineering tools. Here, we present optimized electroporation and polyethyleneglycol (PEG) transformation methods for the model diatom Phaeodactylum tricornutum, enabling delivery of DNA and Cas9 ribonucleoprotein complexes with high efficiency. Transformants are recovered with as little as 1 ng of DNA, and linear or circular episomes as large as 55.6 kb are successfully introduced. The optimized electroporation protocol also reveals an unexpected capability: episomes can be assembled directly in the algal cell through non-homologous or homology-driven repair mechanisms, a process we term diatom in vivo assembly (DIVA). In addition, the PEG approach is adapted to successfully transform Thalassiosira pseudonana, demonstrating the applicability of our methods for engineering other diatom species. These tools could be used to accelerate diatom synthetic biology projects and, therefore, the development of sustainable technologies.

Introduction
Climate change, population expansion, and soil depletion are among several challenges that existentially threaten current agricultural practices. Due to their comparatively rapid growth and higher photosynthetic efficiency, unicellular algae have been proposed as a sustainable agricultural alternative to traditional crops since the 1950s1. Diatoms (Bacillariophyta) are of particular interest for this purpose as they are widely distributed across the world’s oceans2, where they are one of the most photosynthetically productive taxa – accounting for an estimated 20% of total fixed organic carbon annually3.

Phaeodactylum tricornutum is considered to be one of the most characterized model diatom species, having a fully sequenced telomere-to-telomere reference genome4 in addition to molecular tools for DNA transformation5–8, CRISPR-based genome engineering9, protein expression10, cloning of the whole organelle genomes11,12, and more. These characteristics have made P. tricornutum an emerging chassis for diatom biotechnology13 and the subject of a whole-genome synthesis effort14. The latter, termed the Pt-syn1.0 project, has brought together an international consortium of phycologists with the aim of replacing all the native chromosomes in the nuclear, chloroplast, and mitochondrial genomes with synthetic counterparts. To actualize this, and to discover the full potential of diatoms as cell factories, a pipeline for efficiently and rapidly testing different DNA constructs in P. tricornutum must be established.

Existing methods for delivering DNA to P. tricornutum include biolistic bombardment8, electroporation5,7, polyethyleneglycol (PEG) transformation6, and bacterial conjugation6. Biolistics was the first method developed for transforming P. tricornutum and has been used to integrate non-replicative DNA into the nuclear genome8,15, with reported efficiencies of 1 to 100 transformants per 108 cells. Though reasonably efficient, this method requires the use of expensive equipment (i.e., a biolistic particle delivery system) that many labs do not have access to; comparatively, conjugation, electroporation, and PEG transformation can all be conducted with more standard equipment.

Among these, conjugation is the most efficient transformation method for delivering episomes (4 transformants per 104 cells)6, but also the most mutation prone and laborious to execute6,9. Conjugation requires the creation and maintenance of a donor strain of Escherichia coli, which must contain a suitable broad-host-range conjugative plasmid in addition to the episome of interest; complications can arise if these two constructs have incompatible replication machinery16, if the episome is maintained at medium to high copy number6, or if it contains elements that are toxic when expressed in E. coli (e.g., endonucleases)9. These issues can lead to episomal rearrangements or loss over time. As a result, donor strains often require repeated reisolation and plasmid sequencing to confirm construct stability – steps that can add days to weeks, and in some cases months, to a workflow.

The PEG and electroporation transformation methods offer a more direct path to episomal delivery, but each has limitations. The delivery of episomes through PEG transformation remains unutilized due to its low efficiency (≤ 1 transformant per 108 cells) and high variability between experiments6. Although PEG-mediated transformation has not yet been demonstrated in other diatom species, it is routinely applied in the microalga Chlamydomonas reinhardtii, where cell-wall removal followed by PEG-mediated agitation with DNA and glass beads enables efficient nuclear transformation17. Electroporation has a higher transformation efficiency (2.6 to 4.5 transformants per 105 cells)5,7, but has not received wide adoption due to variability in efficiency and issues with reproducibility. Thus, we sought to optimize the PEG and electroporation methods for episome delivery to P. tricornutum with the hopes of developing a more rapid alternative to bacterial conjugation.

In this paper we describe optimized electroporation and PEG transformation protocols for engineering P. tricornutum. These methods are both faster and simpler to execute than bacterial conjugation, as well as more efficient than previously described protocols. We also demonstrate that it is possible to assemble episomes directly in the algal cell through non-homologous repair (NHR) and homology-directed repair (HDR) pathways, opening up avenues for exploring alternative assembly strategies that do not rely on traditional E. coli or S. cerevisiae cloning. Both protocols are suitable for rapid high throughput testing of several DNA constructs simultaneously and, due to their efficiency with low DNA amounts, could be used for the delivery of commercially synthesized episomes directly into the algal cell. Electroporation can also be used for the efficient delivery of Cas9 ribonucleoprotein (RNP) complexes, enabling DNA-free genetic engineering. The protoplasting techniques we establish in P. tricornutum are also used to successfully engineer Thalassiosira pseudonana cells via PEG transformation, demonstrating that these methods can be applied to other diatom species. These tools will be valuable as the Pt-syn1.0 project begins to build and test different synthetic chromosome designs.

Results
Electroporation efficiency increases with enzymatic treatment of cells

Our initial attempts at electroporation using a previously described protocol18 that was adapted from the methods of Zhang and Hu5 yielded no transformants. After several failed attempts, we discovered that adjusting the electroporator capacitance from 25 to 50 µF yielded transformants for both episomes and marker cassettes, though efficiency was highly variable between experiments (Supplementary Data 1, Exp. 1 to 81; Supplementary Fig. 1). We used this method to deliver a linear 11 kb PCR-amplified episome, pPtGE31_ΔPtR (Addgene ID: 236260), to assess whether the construct would integrate into the genome, persist as a linear episome, or be converted into a circular episome within the P. tricornutum nucleus. We were able to recover DNA from these algal transformants in EPI300 E. coli, indicating that the linear episome had been circularized in the algal nucleus following electroporation (Supplementary Fig. 2). Whole-plasmid sequencing of four E. coli colonies confirmed that circularization had occurred via fusion of the construct’s termini.

When this electroporation protocol was repeated using cells that had reached the late stationary phase of growth, we obtained up to 272 transformants per 108 cells with the same 11 kb PCR-amplified episome (Supplementary Data 1, Exp. 82 to 89). This represented an nearly 20-fold increase in efficiency when compared to transformations with early-log phase cultures (Exp. 80 and 81). Stationary phase cultures demonstrated noticeable differences in cell phenotype when compared to early-log phase cultures, with there being a higher abundance protoplasts (i.e., diatoms lacking a cell wall) in the older cultures. P. tricornutum cells can appear in three different morphotypes – fusiform, oval, and triradiate – and can switch morphotypes in response to environmental stimuli19,20. All morphotypes can generate protoplasts if the cell wall is sufficiently weakened. Ultimately, this observation led us to hypothesize that cell morphotype and cell wall integrity may influence electroporation transformation efficiency. Notably, a spheroplasted cell has only partial cell wall degradation and may not appear phenotypically different from a wild-type cell, whereas a fully protoplasted cell lacks a cell wall entirely and adopts a characteristically spherical morphology – provided it does not burst from osmotic or mechanical stressors.

Cell morphotype cannot be readily controlled in P. tricornutum cultures as this species displays a high degree of phenotypic plasticity; however, agar-plated cultures of CCAP1055/1 tend to have a higher proportion of oval-type cells (~20–30% oval, ~70–80% fusiform) compared to liquid cultures (<10% oval, >90% fusiform). Thus, to test this hypothesis, we spheroplasted early-log phase P. tricornutum cells harvested from agar plates. Plate-derived cells were spheroplasted using the enzyme alcalase, a serine protease that has been shown to degrade the proteinaceous components of the P. tricornutum cell wall21. We observed that 100 µl of alcalase at an activity of approximately 3 Anson units per ml (i.e., 300 mAnson units) was enough to fully protoplast 3 × 108 plate-derived cells when resuspended in 375 mM D-sorbitol (Fig. 1a, b). The protoplasted cells were very fragile and prone to rupture from osmotic pressure or physical damage; treatment of 3 × 108 cells with 10 µl of alcalase (i.e., 30 mAnson units) was enough to protoplast some cells, whilst leaving others with partially degraded cell walls (i.e., spheroplasts; Supplementary Fig. 3). Plate-derived cells (3 × 108) were treated with 0.3 mAnson units, 3 mAnson units, and 30 mAnson units of alcalase prior to the washing steps of the electroporation protocol. Transformation efficiency increased in all cases (Fig. 1c) when electroporating 500 ng of the same 11 kb PCR-amplified episome as before (Fig. 1d), with the 3 mAnson treatment demonstrating the highest increase in efficiency. Based on this result, we went forward with using 1 mAnson unit of alcalase per 108 cells when spheroplasting P. tricornutum for electroporation.

Electroporation efficiency of cells grown in liquid and solid media

We tested the electroporation efficiency of P. tricornutum cells grown in liquid media and on agar plates, with and without alcalase treatment across five biological replicates (Fig. 1e) using 500 ng of the same PCR-amplified 11-kb episome as before. Though transformation efficiency varied between experiments, plate-derived alcalase-treated cells were significantly more efficient (~1.2 transformants per 104 cells) than untreated plate-derived cells (~6.0 transformants per 107 cells), untreated liquid-derived cells (~4.2 transformants per 107 cells), and treated liquid-derived cells (~1.6 transformants per 106 cells, Supplementary Table 1). Statistical significance was determined using a Kruskal-Wallis test (p = 0.0094) followed by pairwise Wilcoxon tests with Bonferroni correction. Plate-derived, alcalase-treated cells yielded significantly more colonies than all other groups (p < 0.05), whereas no significant differences were observed among the remaining comparisons.

Based on this, all proceeding electroporations were conducted with cells derived from plates and spheroplasted with 1 mAnson unit of alcalase per 108 cells. This optimized method was compared with the original and adjusted electroporation protocols to demonstrate the impact that changing the capacitance parameter and spheroplasting have on electroporation efficiency (Supplementary Fig. 1).

Transformation efficiency with varying amounts of DNA

Our prior transformations were all conducted with ~500 ng of a PCR-amplified 11-kb plasmid, pPtGE31_ΔPtR (Fig. 1d). We were interested to see how transformation efficiency varied when using differing amounts of this construct, and most importantly, what the minimum amount of DNA for transformation is. We electroporated as little as 1 ng and up to 1 µg of PCR-amplified pPtGE31_ΔPtR across four biological replicates (Fig. 1f, data summarized in Supplementary Table 2). In all but one experiment, we were able to recover transformants when using just 1 ng of DNA during electroporation. Increasing the amount of DNA to 10 ng routinely gave rise to hundreds of transformants, with there being an average of 372 CFUs per reaction. Interestingly, there was only a small change in transformation efficiency when the amount of DNA was doubled from 500 ng to 1 µg, suggesting that 500 ng of linear pPtGE31_ΔPtR was enough to nearly saturate the electroporation reaction.

Transforming linear and circular episomes to the algal nucleus

Our past explorations had relied upon the use of linear PCR-amplified DNA with blunt, non-phosphorylated termini. To test the efficiency of different structural forms of DNA, we prepared other variations of pPtGE31_ΔPtR (Fig. 2a). The termini of the linear constructs varied in position along the episome and/or the properties of the termini (i.e., blunt or sticky ends, with or without 5′ phosphorylation). These constructs were adjusted to be equimolar prior to conducting electroporation in P. tricornutum. We found that there were nearly double the amount of CFUs when transforming any structural form of linear DNA compared to the circular episome (Fig. 2b).

We had also demonstrated that electroporated linear pPtGE31_ΔPtR gets circularized in the P. tricornutum nucleus, but we had not yet performed comprehensive screening to determine if any indels were accruing during this repair process. To explore what occurs during termini fusion, ten colonies were repatched from the transformations with circular and PCR-amplified pPtGE31_ΔPtR for additional screening. Colonies were screened using a set of primers that spans where the termini are expected to recombine for the PCR-amplified construct, as well as a region of the episome backbone (Fig. 2c). Colonies transformed with circular DNA did not demonstrate any unexpected insertions or deletions in the screened regions (Fig. 2d), as expected. Whole plasmid sequencing of circular colonies 1 and 2 confirmed that the episomes had not accrued any mutations or indels during transformation.

Colonies transformed with PCR-amplified DNA appeared to have indels in the region where termini fusion occurs but not in the intact backbone region (Fig. 2e). We conducted Sanger sequencing of the termini region for colonies 1, 2, and 6 to 10, which confirmed that indels of varying sizes had accrued in this region for every colony screened (Supplementary Fig. 4). The absence of the 486 bp band in lanes 3, 4, and 5 is most likely due to DNA degradation or deletions occurring at the episomal termini, either before (e.g., exonuclease activity) or during the circularization process. Whole plasmid sequencing of linear colonies 1 and 8 further confirmed the presence of indels in the termini region where the episome was being fused together, whereas all other regions of the episome had remained unchanged. Interestingly, the ~500-bp insertions in colonies 2, 8, and 10 were not complementary to any regions of the episome or algal genome. When these insertions were queried using BLASTn, they aligned to various regions of the salmon and/or trout genome accessions.

Single-stranded salmon sperm (ssss) DNA is used as a carrier during electroporation, and to our surprise, we had discovered that it could be captured during NHR of the episome termini. We attempted electroporation without ssssDNA to see if this would impact transformation efficiency, and it resulted in a 15-fold reduction in the number of CFUs (Supplementary Fig. 5), highlighting its importance as an additive for efficient transformation.

Evaluation of genomic integration

We performed a plasmid/episome loss experiment by serially passaging four pPtGE31_ΔPtR transformants in non-selective media to check for genomic integration. The transformants chosen correspond to algal colonies 1 and 2 in Figs. 2d and 1 and 8 in Fig. 2e. After seven passages in liquid media with or without nourseothricin selection, serial dilutions of the cultures were then plated onto solid non-selective media to obtain single colonies. For each transformant, 100 single colonies were then re-patched onto plates supplemented with nourseothricin. If genomic integration of the episome had occurred, we would expect a high proportion of the single colonies to retain resistance. Conversely, if the episomes are maintained extrachromosomally, they should be lost over successive generations without selective pressure, and thus only a low proportion of colonies would retain resistance. As expected, only 2–7% of screened colonies retained resistance (Supplementary Table 3), demonstrating that genomic integration had not occurred in the assayed transformants.

Electroporation and recovery of a large plasmid

It has been demonstrated in other organisms that as episome size increases, electroporation efficiency decreases22,23. We sought to test the limits of electroporation in P. tricornutum by attempting transformation with pSC5, a 55.6 kb broad-host-range conjugative episome (Fig. 3c, Addgene ID: 188602)24.

Equimolar amounts of circular and linear pSC5 were prepared along with PCR-amplified pPtGE31_ΔPtR (Fig. 3a). Electroporation with either form of pSC5 gave rise to transformants, though the number of CFUs was orders of magnitude lower than electroporation with a linear 11 kb episome (Fig. 3b), as anticipated. To determine if the complete 55.6 kb episome had been successfully electroporated into P. tricornutum, we passaged four colonies from each of the transformations with circular and linear pSC5. DNA was isolated from the algal colonies and electroporated into E. coli. Here, if the E. coli transformants contain an intact version of pSC5, the episome will facilitate conjugation and self-mobilization into a recipient strain harboring pSAP12 (Addgene ID: 206429), a 10 kb plasmid that contains a chloramphenicol resistance marker (Fig. 3d). Successful transconjugants will harbor pSC5 and pSAP, therefore carrying resistance to both gentamycin and chloramphenicol.

We performed conjugation using a pool of E. coli colonies for each transformation (Fig. 3e; C1–C4 are derived from circular pSC5 algal transformants, L1 to L4 are derived from linear pSC5 algal transformants). Conjugation was successful for three out of four (circular pSC5) and four out of four (linear pSC5) of the E. coli pools tested, demonstrating that intact pSC5 had been successfully electroporated and maintained in the majority of screened P. tricornutum transformants (Fig. 3e).

Electroporation and recombination of multiple DNA fragments

Assembly via a non-homologous repair pathway. The surprising discovery of salmon DNA inserted into the termini region of PCR-amplified pPtGE31_ΔPtR led us to hypothesize that P. tricornutum could assemble multiple DNA fragments into a single episome through NHR. To test this, we created an episome, pPtGE31_ShBle (Addgene ID: 236261), that contains the NAT and ShBle markers conferring resistance to nourseothricin and zeocin, respectively. The episome pPtGE31_ShBle was then amplified as two non-overlapping fragments, each containing one of the algal selective markers. The fragments were adjusted to be equimolar prior to electroporation into P. tricornutum (Fig. 4a).

Cells transformed with both fragments simultaneously were plated on nourseothricin- or zeocin-containing plates initially (Supplementary Fig. 6). Out of the 116 colonies passaged post-electroporation, 94 demonstrated double resistance to both nourseothricin and zeocin (Supplementary Table 4). Twenty transformants, ten derived from each initial transformation plate, were passaged onto double-selection media, then DNA isolation and PCR screens were performed. These colonies are denoted with an N- or Z- to depict which initial selection plate they were derived from (N: nourseothricin, Z: zeocin).

Two orientations are expected when the fragments amplified from pPtGE31_ShBle are re-assembled (Fig. 4b). We designed a PCR screen that would not only assess if both fragments had been recombined, but could also determine the orientation of the fragments. When screening algal transformants, we only used the primer set spanning junction B as junction A contains primers that bind to the FcpD promoter and FcpA terminator, which are present in both the episome and algal genome. Of the twenty colonies screened, nine had amplification across junction B, with only five colonies demonstrating the expected amplicon sizes (Supplementary Fig. 7). The other four colonies either demonstrated multiple amplicons, some of which were the expected sizes, and/or amplicons that were larger than expected. Based on this initial screen, DNA from colonies N-C6, N-C9, N-C10, Z-C2, Z-C4, and Z-C5 was transformed into E. coli for further screening. Three E. coli transformants were assessed per algal colony using the primer sets spanning junctions A and B (Fig. 4c, d). Interestingly, the E. coli transformants derived from algal colonies N-C6, N-C10, and Z-C2 demonstrated different banding patterns, suggesting that these algal colonies likely had a pool of assembled episomes. All E. coli transformants from algal colony N-C9 depicted the expected banding pattern for orientation 2, whereas all E. coli transformants from algal colony Z-C5 depicted the expected banding pattern for orientation 1. DNA from E. coli colonies N-C6-E1/E2, N-C9-E1, N-C10-E1/E2, Z-C2-E1/E2/E2, Z-C4-E1, and Z-C5-E1 were sent for whole-plasmid sequencing to validate if assembly had occurred.

The sequencing results demonstrated that recombination of fragments 1 and 2 had occurred in all assessed P. tricornutum transformants (Supplementary Fig. 8). In the majority of assembled episomes, multiple partial copies of fragments 1 and 2 had recombined. The most excessive recombination appeared E. coli colony Z-C2-E2, which harbored a 41.5 kb episome containing eight partial or whole copies of both fragments 1 and 2. Only E. coli colonies N-C9-E1, N-C10-E1, Z-C2-E3, and Z-C5-E1 contained the expected assembled episomes. Sequencing of E. coli transformants derived from the same algal colony revealed that algal colonies N-C6, N-C10, and Z-C2 possessed a pool of assembled episomes, as we had hypothesized.

Assembly via a homology-directed repair pathway. Though it is possible to assemble episomes via the NHR pathway, screening and sequencing demonstrated that the majority of P. tricornutum transformants contained undesirable assemblies of the non-overlapping fragments. To be able to reliably assemble episomes in the algal nucleus, an alternative strategy is needed. We sought to test whether overlapping fragments could be simultaneously transformed and assembled through the HDR repair pathway. It was hypothesized that the overlapping sequences between fragments would enable site-specific recombination at the termini, thereby creating episomes of an expected size and directionality.

To test this hypothesis, we split the episome pPtGE31_ShBle into two, three, or four fragments that overlapped with each other by ~160 bp (Fig. 5a). This length was chosen somewhat arbitrarily as a starting point, as we were uncertain about the minimum or maximum overlap size needed for successful HDR. Fragments were generated by PCR amplification using PtGE31_ΔPtR (Addgene ID: 236260) or pPtGE31_ShBle (Addgene ID: 236261) as template DNA (primers listed in Supplementary Data 2). In all assembly designs, one of the overlaps was positioned within the open reading frame (ORF) for nourseothricin N-acetyl transferase (NAT), which confers nourseothricin resistance. Thus, when the assembly fragments are electroporated, nourseothricin-resistant transformants should only appear if homologous recombination has occurred within the NAT ORF or if there is carry-over of the PCR template DNA. Primers were designed to screen the other overlapping termini for successful recombination (Fig. 5a, bottom boxes).

We first tested the two-piece HDR assembly by electroporating fragments 1 and 2 individually or simultaneously (Supplementary Fig. 5b). When electroporated individually, fragments 1 and 2 gave rise to 8 and 2 CFUs, respectively, indicating that carry-over of the PCR template DNA had occurred. However, when the fragments were electroporated simultaneously, there were more than 1800 CFUs – far more than what would be anticipated by carry-over alone. Ten algal colonies were selected for screening of the other fragment junction (Supplementary Fig. 9a), all of which demonstrated the expected 300 bp band, suggesting that successful homologous recombination had occurred. Episomes from algal colonies C1, C2, and C3 were recovered in E. coli. Three E. coli colonies were screened per initial algal colony, all of which demonstrated the correctly sized band (Supplementary Fig. 9b). To confirm if the fragments had been correctly recombined, episomes from E. coli colonies C1-E1, C2-E1, and C3-E1 were sent for whole plasmid sequencing. This demonstrated that colonies C1-E1 and C3-E1 contained the correctly assembled episome (Supplementary Fig. 10), but the episome recovered from C2-E1 had two partial copies of fragment 1.

The same methodological approach was used to electroporate three overlapping fragments simultaneously (Fig. 5c). Electroporation of fragments 1 and 2 or 2 and 3 gave rise to 38 and 47 CFUs respectively, whereas electroporation of all three fragments gave rise to ~6500 CFUs. Ten algal colonies were screened for the junctions between F1/F2 or F2/F3. All colonies contained the expected bands except for C1 and C2, which lacked an amplicon for the F1/F2 junction and demonstrated a weak amplicon for the F2/F3 junction (Supplementary Fig. 9c). We anticipated that assembly may have failed in these colonies, but to further investigate this, we attempted to recover the algal episomes from C1 and C2 in E. coli, along with the positively screened episomes from C3 and C4. We were able to recover E. coli transformants for all the tested episomes and decided to screen two CFUs per initial algal colony, all of which demonstrated correctly sized amplicons for both junctions (Supplementary Fig. 9d). Episomes isolated from E. coli colonies C1-E1, C2-E1, C3-E1, and C4-E1 were sent for whole plasmid sequencing, with only C1-E1 deviating from the expected assembly (Supplementary Fig. 10).

To further test the limits of the HDR assembly pathway, we attempted electroporation of four overlapping fragments simultaneously. Electroporation of only three of the four fragments (i.e., F1+F2+F3 or F2+F3+F4) gave rise to 85 and 56 transformants, demonstrating the carry-over of template DNA once again (Fig. 5d). However, when all four fragments were combined and transformed simultaneously, there was a >20-fold increase in the number of CFUs. Ten algal colonies were screened across the remaining three junctions, and all but C1 and C7 demonstrated the expected amplicons (Supplementary Fig. 9e). Colonies C1 and C7 specifically lacked the 273 bp amplicon for the junction between F1/F2. Algal episomes from C1, C2, C3, and C7 were transformed into E. coli; however, C7 did not give rise to any bacterial transformants, suggesting that incomplete assembly had occurred. Two E. coli colonies were screened for the episomes derived from C1, C2, and C3, all of which demonstrated the expected amplicons for the three junctions (Supplementary Fig. 9f). Whole plasmid sequencing of C1-E1, C2-E1, and C3-E1 revealed that correct assembly had occurred in algal colonies C1 and C3 (Supplementary Fig. 10), but C2 contained multiple partial copies of fragments 1 and 4. In total, 26 of the 30 algal colonies screened across the various assemblies demonstrated that recombination had successfully occurred across the tested junctions (Supplementary Fig. 10). From the episomes recovered in E. coli, 7 out of 10 had the correctly assembled episomes when fully sequenced. This demonstrates that it is both feasible and reliable to assemble episomes through the HDR pathway in P. tricornutum.

Exploring the limits and applications of HDR assembly

The discovery of DIVA prompted us to further investigate the practical boundaries and potential applications of this assembly method. We first sought to determine the size of overlaps required to direct HDR. Our earlier assemblies relied on ~160 bp overlaps, but in Saccharomyces cerevisiae, as little as 20 bp is sufficient for recombination25. To test whether this was also true in P. tricornutum, we PCR-amplified NAT using primers that generated 0, 20, or 40 bp of overlap with the FcpD and FcpA promoter/terminator pair (Supplementary Fig. 11a). We then PCR-amplified pPtGE30 (Addgene ID: 236256) in two fragments, positioning the termini such that the NAT constructs would assemble into the Sh Ble locus, which uses the same promoter/terminator elements.

The NAT constructs were electroporated both individually and with the episomal fragments (Supplementary Fig. 11b). Electroporation of NAT-0, NAT-20, and NAT-40 alone confirmed that the marker with up to 40 bp of promoter/terminator sequences does not confer nourseothricin resistance. The NAT-0 assembly produced no transformants, whereas both the NAT-20 and NAT-40 assemblies yielded ~100 transformants. These results indicate that overlaps as short as 20 bp are sufficient for HDR-mediated assembly.

We next examined whether de novo synthesized constructs could be inserted directly into episomes in P. tricornutum. We commercially synthesized two constructs: (i) CytB-eGFP (833 bp), which carries the CytB mitochondrial targeting peptide identified by TargetP (2.0)26, and (ii) eGFP-T2A-NAT (1025 bp), in which eGFP is linked to residues 2 to 63 of the NAT protein via the T2A ribosomal skip sequence27. The CytB-eGFP construct was designed to insert into the Sh Ble locus of pPtGE31_ShBle (Supplementary Fig. S11c), whereas the eGFP-T2A-NAT construct was designed to insert into a PCR-amplified version of pPtGE31_ΔPtR containing an incomplete NAT cassette (residues 47–188; Supplementary Fig. 11d). In both designs, the synthesized constructs included 50 bp of overlaps at each terminus to their respective insertion sites.

For the eGFP-T2A-NAT design, nourseothricin-resistant colonies should only arise if correct assembly restores the complete NAT gene or if there is unintended carry-over of the template episome. This reduces the likelihood of false positive transformants. Both de novo assembly designs yielded at least one hundred colonies (Supplementary Fig. 11e, f).

Ten nourseothricin-resistant colonies from each assembly were passaged and screened for GFP expression by confocal fluorescence microscopy. Of the CytB-eGFP transformants, 4/10 showed moderate mitochondrial eGFP localization, while all 10/10 of the eGFP-T2A-NAT transformants exhibited strong cytosolic eGFP expression (representative images in Supplementary Fig. 12). Three colonies per design were subsequently recovered in E. coli and sequenced (C3, C5, C7 for CytB-eGFP; C1, C2, C3 for eGFP-T2A-NAT). All recovered episomes contained the corresponding de novo construct; however, only one CytB-eGFP episome (C5-E1) and one eGFP-T2A-NAT episome (C1-E1) displayed the expected assembly configuration. The remaining episomes showed unexpected recombination events, including combinations of multiple episomal fragments and, in some cases, insertion of multiple copies of the de novo construct or unaligned DNA.

Delivery of RNPs via the optimized electroporation protocol

RNP complexes can be delivered to P. tricornutum through biolistic bombardment28, providing a means for DNA-free genome engineering. We anticipated that it would also be possible to electroporate RNPs into this species, but this had yet to be reported in any diatoms. As a proof-of-concept, we replicated the methodology Serif et al. had used to deliver RNPs28 using our optimized electroporation protocol in lieu of biolistics.

Serif et al. had previously shown that knock-out of the P. tricornutum Adenine Phosphoribosyl Transferase (PtAPT) gene confers resistance to the cytotoxin 2-fluoroadenine (2-FA) at a dose of 10 µM28. PtAPT functions in the adenine salvage pathway, catalysing the transfer of phosphoribosyl pyrophosphate (PRPP) to adenine to form adenosine monophosphate (AMP); in the presence of 2-FA, PtAPT instead converts 2-FA into 2-fluoro-AMP, which is subsequently phosphorylated to form toxic fluorinated nucleotides28.

Chemicals can vary based on the manufacturer and/or batch; thus, when setting forth on this series of experiments, we first tested the resistance of wild-type P. tricornutum (CCAP1055/1) cultured on ½-salt L1 plates supplemented with adenine (5 µg/ml) and our batch of 2-FA (0, 2.5, 5, 10, 15, 20 µM). To our surprise, there was an absence of growth on the 0 µM 2-FA plate, suggesting that adenine alone posed a toxicity to wild-type P. tricornutum cells. We tested this theory by plating wild-type cells onto ½-salt L1 plates supplemented with varying levels of adenine (0, 1, 2.5, 5 µg/ml; Figure S13). The growth of wild-type algae was negligible after one-week on the 5 µg/ml adenine plate, but relatively unimpacted on the 0.5 and 1 µg/ml plates. This interesting revelation led us to hypothesize that adenine supplementation could negatively impact ΔPtAPT transformants as well, so when conducting our RNP electroporation experiments, we plated transformants on 2-FA plates with and without adenine supplementation.

The PtAPT locus was targeted using two guide RNAs (gRNAs), each of which allowed for biallelic knock-out; we opted to use the first gRNA (gAPT1) for our proof-of-concept experiments as it had generated 11 of the 16 2-FA-resistant transformants in the singleplex experiments by Serif et al.28. HiFi Cas9 and a single guide RNA for APT1 (sgAPT1) were complexed according to the manufacturer’s guidelines (Fig. 6a). Then, the RNP complex was combined with ssssDNA and electroporated into spheroplasted P. tricornutum cells. We opted to include carrier DNA when performing the RNP experiments as ssssDNA is thought to bind to and neutralize the positive charge of the cell membrane during electroporation, making it easier for other charged biomolecules to pass through29.

Transformants were recovered for one day before plating one-fifth of the reaction across ½ salt L1 plates supplemented with 10 µM or 5 µM 2-FA, with or without 5 µg/ml adenine (Supplementary Fig. 14). This pilot experiment generated thousands of 2-FA-resistant transformants across the plates that had been supplemented with adenine, but only a handful across the plates without supplemental adenine. This was an unexpected discovery given our previous observations. Ten 2-FA-resistant colonies were passaged for further screening at the PtAPT loci to determine if Cas9-mediated engineering had occurred.

We used the primer sequences reported by Serif et al.28 (Supplementary Data 2) to screen the PtAPT alleles using PCR. When using primers that bind to both alleles, amplicons of varying sizes were visible across the 2-FA resistant transformants, most of which differed from that of the wild-type strain (Fig. 6b). Likewise, amplicons of varying sizes were visible when using primers specific to allele 1 (Figs. 6c) and 2 (Fig. 6d). This suggested that Cas9-mediated engineering had occurred, but to corroborate our findings, amplicons generated with allele-specific primers were sent for Sanger sequencing.

Alignment of the sequenced amplicons to the wild-type sequence for alleles 1 and 2 confirmed the presence of indels in all 12 alleles that could be analysed (Fig. 6e). We were able to generate sequencing data of both alleles for three of the transformants (C1, C6, and C7), with one clone (C1) demonstrating a homozygous 12 bp deletion across both alleles. Interestingly, there were large (>50 bp) insertions in four of the sequenced alleles. Querying the insertion sequences in NCBI BLAST revealed that carrier DNA had integrated into these loci during the double-strand break repair process. The alignments for C6 and C10 contained two and four non-contiguous regions from the salmon and/or trout genomes, demonstrating that multiple shorn pieces of carrier DNA had been integrated as opposed to a single large fragment.

Cotransformation of an RNP complex and episome

With the discovery that RNP complexes can be electroporated into P. tricornutum, we next sought to see if both an RNP and episome could be co-transformed simultaneously, and if so, what the rate of co-transformation is. The aim of this experiment was to develop a protocol that could significantly improve screening for gene deletions that do not produce an obvious phenotype. To test this, the PCR-amplified episome pPtGE31_ΔPtR was combined with the Cas9:sgAPT1 complex and electroporated into spheroplasted P. tricornutum cells. Transformants were first plated on nourseothricin or 2-FA selective plates, and then later repatched onto the alternative selection type to determine if co-transformation had occurred.

Our earlier RNP transformation experiment (Supplementary Fig. 14) had demonstrated that adenine supplementation is important when selecting for PtAPT knock-out colonies post electroporation; however, adenine is also seemingly toxic to untransformed P. tricornutum cells (Supplementary Fig. 13). Thus, when co-transforming pPtGE31_ΔPtR and Cas9:sgAPT1, the electroporated cells were plated on two different selection plates: (1) ½-salt L1 plates supplemented with 5 µg/ml adenine and 10 µM 2-FA, and (2) ½-salt L1 plates supplemented with 1 µg/ml adenine and 100 µg/ml nourseothricin. The lower adenine concentration was chosen to allow for the growth of co-transformed colonies whilst not unintentionally selecting against colonies that had only uptaken the episome.

After two weeks of growth on nourseothricin or 2-FA selection, 50 co-transformed colonies were repatched onto plates with the same selection type and grown for 1 week. Then, the colonies were repatched onto the alternative selection type to determine if any had double-resistance. Of the 100 screened colonies, 13 demonstrated double-resistance to nourseothricin and 2-FA (Supplementary Table 5), indicating co-transformation had successfully occurred at a rate of ~10%.

Revisiting the PEG transformation method

A polyethyleneglycol (PEG) transformation method for P. tricornutum with an efficiency of ≤ 1 transformant per 108 cells was previously described by Karas et al.6. For this method, algal cells were protoplasted using a combination of zymolyase, lysozyme, and hemicellulose ahead of PEG treatment. We revisited this method using alcalase as the sole enzyme for cell wall removal, with the intention of providing a simple and equipment-independent alternative to electroporation that is particularly useful for labs or species where this method remains a practical barrier.

We followed the method described by Karas et al.6, substituting the aforementioned enzyme cocktail with 10, 100, or 1000 µl of alcalase (i.e., 30 mAnson, 300 mAnson, and 3 Anson, respectively) for protoplasting ~3 × 108 cells. The 3 Anson treatment was potent enough to protoplast the majority (i.e., >80%) of the cells. As well, the protoplasts were transformed with 500 ng of linear pPtGE31_ΔPtR as opposed to 25 µg of a circular episome, the amount originally used by Karas et al., to be able to compare the efficiency of transformation with our electroporation experiments. Treatment with 3 Anson of alcalase generated approximately 160 transformants per 3 × 108 cells, an efficiency of ~4 transformants per 107 cells. The other two treatments yielded 0 transformants (30 mAnson of alcalase) and approximately 6 transformants (300 mAnson alcalase), so all future PEG transformations were conducted with 3 Anson of alcalase per 3–5 × 108 cells. This suggests that complete cell wall removal is paramount to efficient PEG transformation.

We then sought to test other parameters of the PEG method to further optimize the transformation efficiency. First, we began to include 40 µg of carrier DNA per PEG reaction as this had led to a ~15-fold increase in transformation efficiency during electroporation. Then, we tested various concentrations of PEG-8000 and found that a range from 20 to 30% (w/v) can be used (Fig. S15a). Going forward with 20% PEG, we then tested if the duration of protoplasting, inclusion of carrier DNA, and length and/or temperature of incubation would alter transformation efficiency (Fig. S15b). As anticipated, the inclusion of carrier DNA increased transformation efficiency >10-fold (Supplementary Fig. 15c). Longer protoplasting and incubation times also appeared to increase transformation efficiency to ~6 transformants per 106 cells (Supplementary Fig. 15c).

The optimized PEG transformation protocol was then used to transform 1 µg of pSC5 into protoplasted P. tricornutum cells, generating ~60 transformants (Supplementary Fig. 15d). DNA were isolated from four algal transformants and recovered in EPI300 E. coli. Bacterial transformants were then assessed for conjugation in the same manner as before. All recovered episomes demonstrated the ability to conjugate (Supplementary Fig. 15e), verifying that pSC5 had been intactly transformed into P. tricornutum through the PEG method.

PEG transformation to T. pseudonana

Electroporation parameters can vary drastically between species and are often finnicky to troubleshoot or optimize. To expand the applications of our alcalase-based transformation methods, we sought to adapt the optimized PEG transformation protocol for T. pseudonana, a model diatom species that possesses a more typical silicified frustule. Given the differences in cell wall composition between these species, T. pseudonana cells were subjected to varying protoplasting conditions (Supplementary Fig. 16a) before PEG treatment. The circular episome pBIG1 (8.2 kb) conferring nourseothricin resistance and carrying eGFP was used in these experiments. All protoplasting conditions led to the emergence of colonies on the nourseothricin selection plates, with the harsher protoplasting conditions (i.e., ≥ 1.5 Anson alcalase) yielding more growth.

Transformants from the 500 µl alcalase treatment plate (i.e., 1.5 Anson, 40 min) were pooled and subjected to fluorescence microscopy to assess if eGFP was being expressed, as this would indicate successful episome transfer. Both autofluorescence and eGFP signals were visible in the transformed cells (Supplementary Fig. 17), demonstrating that episome transfer had occurred.

To further characterize if protoplasting was truly occurring, T. pseudonana cells were incubated with 10 or 1000 µl of alcalase (i.e., 30 mAnson and 3 Anson, respectively) for 40 min (Supplementary Fig. 16b and c, respectively) and then imaged via microscopy. Based on these microscopy images, cells from the 10 µl treatment (i.e., 30 mAnson) showed a protoplasting rate of ~4% (1/28 cells), whereas cells from the 1000 µl treatment (i.e., 3 Anson) demonstrated a protoplasting rate of ~50% (13/20 cells). This finding underscores that alcalase can be used to degrade the cell wall of diatoms harboring silica frustules.

Methods
Microbial strains and growth conditions
P. tricornutum (Culture Collection of Algae and Protozoa, CCAP1055/1) was cultured in liquid L1 media or on ½-salt L1 1% agar plates (w/v) in a growth chamber set to 18°C and a light/dark cycle of 16/8 h. Cultures were irradiated with white LED lights (Mars Hydro, model: TSL2000) set to photon flux densities of ~75 μE m−2 s−1 (liquid cultures) or ~55 μE m−2 s−1 (agar plate cultures). Agar plates were placed in clear polystyrene trays to reduce desiccation. L1 media was prepared as previously described and lacks silica6.

In brief, liquid L1 medium was prepared by combining 1 L aquil salts (synthetic seawater), 2 ml NP stock, 1 ml L1 trace metals stock, and 0.5 ml f/2 vitamin solution. Once combined, 2 M NaOH was added until a pH of ~8.0 was achieved and the solution was sterilized through a 0.2-µm filter. For solid medium, sterile liquid L1 medium was mixed 1:1 with autoclaved 2% (w/v) Bacto-agar (all equilibrated to ~60°C) and poured into petri dishes.

Aquil salts were prepared by mixing equal volumes of separate 2X anhydrous and hydrous salt solutions. The 2X anhydrous salt solution was prepared by dissolving the following in 500 ml ddH2O: NaCl (24.5 g), Na2SO4 (4.09 g), KCl (0.7 g), NaHCO3 (0.2 g), KBr (0.1 g), H3BO3 (0.03 g or 3 ml of a 10 mg ml−1 stock), and NaF (0.003 g or 300 µl of a 10 mg ml-1 stock). The 2X hydrous salt solution was prepared by dissolving MgCl2·6H2O (11.1 g) and CaCl2·2H2O (1.54 g) in 500 ml ddH2O.

The NP stock solution was prepared at 100X by dissolving NaNO3 (37.5 g per 100 ml) and NaH2PO4·H2O (2.5 g per 100 ml) in ddH2O. The trace metal stock was prepared at 1000X by dissolving the following in 1 L of ddH2O: FeCl3·6H2O (3.15 g) and Na2EDTA·2H2O (4.36 g), followed by addition of the following components from concentrated stocks: CuSO4·5H2O (9.8 g L-1), 0.25 ml; Na2MoO4·2H2O (6.3 g L-1), 3.0 ml; ZnSO4·7H2O (22.0 g L-1), 1.0 ml; CoCl2·6H2O (10.0 g L-1), 1.0 ml; MnCl2·4H2O (180.0 g L-1), 1.0 ml; H2SeO3 (1.3 g L-1), 1.0 ml; NiSO4·6H2O (2.7 g L-1), 1.0 ml; Na3VO4 (1.84 g L-1), 1.0 ml; and K2CrO4 (1.94 g L-1), 1.0 ml. Lastly, the vitamin stock was prepared at 2000X by adding thiamine-HCl (200 mg L-1), biotin (10 ml L-1 of a 0.1 g L-1 stock), and cyanocobalamin (1 ml L-1 of a 1 g L-1 stock).

T. pseudonana (Bigelow National Center for Marine Algae and Microbiota, catalog number: CCMP1335) was cultured in liquid F/2 media in a growth chamber set to 20°C and 24 hour white light at a photon flux density of 50 μE m−2 s−1.

S. cerevisiae strain VL6-48 (American Type Culture Collection [ATCC], catalog number: MYA-3666) was grown in 2X YPAD media or plated on complete minimal media lacking histidine with 1% agar (w/v) and 1 M D-sorbitol. Yeast cultures were incubated at 30°C, with liquid cultures placed on a shaker set to 225 rpm. Agar plates were sealed with parafilm to reduce desiccation. Dropout media were prepared according to the manufacturer specifications; in brief, a minimal base media (Takara, catalog number: 630411) was combined with a -HIS dropout supplement (Takara, catalog number: 630415) in ddH2O. Once combined, 2 M NaOH was added until a pH of ~5.6 was reached. YPAD media was sterilized through a 0.2-µm filter; dropout media was sterilized by autoclaving.

E. coli strain EPI300 (LGC Biosearch Technologies, Lucigen, catalog number: EC300110) was grown in lysogeny broth (Miller formula) with or without 1% agar (w/v) and supplemented with 15 µg/ml chloramphenicol and/or 40 µg/ml gentamycin and/or 100 µg/ml L-arabinose. Cultures were placed in an incubator set to 37°C, with liquid cultures placed on a shaker set to 225 rotations per minute (rpm).

PCR amplification
Episomes and/or fragments for transformation and/or assembly were amplified with GXL polymerase (Takara, catalog number: R050A) using the rapid PCR protocol (all primers are listed in Supplementary Data 2). Of note, we used both phosphorylated and non-phosphorylated primers when amplifying pPtGE31_ΔPtR as a linear episome. PCR reactions were conducted with a total of 25 to 35 cycles depending on amplification efficiency. Fragments that were amplified from an episome template were treated with 10 units (0.5 μl) of DpnI (New England Biolabs, catalog number: R0176) and incubated at 37°C for 30 min before deactivation at 80°C for 20 min. After deactivation, fragments were column-purified using the PureLink PCR Purification Kit (Invitrogen, catalog number: K310001) and eluted with ddH2O. Where possible, primers were designed using Primer3 (version 4.1.0).

PCR screening of algal and E. coli transformants
DNA isolated from algal and E. coli transformants was screened for the presence of specific episomal regions using a PCR assay. Here, 1 μl of DNA isolated through alkaline lysis was used as template in a 10 μl SuperPlex PCR reaction (Takara, catalog number: 638543). For all algal transformants, episomes were analzyed with a single primer set at a time as there was rampant off-target binding when multiple primer pairs were used simultaneously (all primers are listed in Supplementary Data 2). We were able to successfully conduct multiplex PCR using E. coli template DNA; however, when screening the NHR assembled episomes, we chose to perform singleplex screens due to the unexpected recombination of the fragments. Reactions were carried out according to the manufacturer guidelines for a total of 30 cycles.

DNA concentrations were measured by fluorometry using the DeNovix dsDNA Broad Range Assay (FroggaBio, catalog number: DSDNA-BROAD-EVAL) and purity ratios were assessed using a Nanodrop spectrophotometer. For the experiments shown in Figs. 2 to 4, samples were adjusted to be equimolar prior to conducting electroporation.

DNA isolation
Episomes were isolated from P. tricornutum, S. cerevisiae, and E. coli using an alkaline lysis protocol that was previously described in Karas et al.6, with species-specific modifications detailed below. Briefly, cell pellets were resuspended in 250 µl of P1 (Qiagen, catalog number: 19051), with or without additional protoplasting enzymes as indicated below, and lysed with 250 µl P2 buffer (Qiagen, catalog number: 19052). Samples were mixed by 5–10 inversions and incubated for 1 min before neutralization with 250 µL P3 buffer (Qiagen, catalog number: 19053). Lysates were centrifuged at 16,000 x G for 10 min, and ~700 µl of supernatant was transferred to a fresh tube. DNA was precipitated from the lysate by adding 750 µl ice-cold 100% isopropanol, which was then centrifuged at 16,000 × g for 10 min. The DNA pellets were washed with 500 µl ice-cold 70% ethanol and centrifuged for 5 min. Pellets were air-dried and resuspended in 30–50 µl sterile ddH2O.

Steps 1 to 2 for E. coli: (1) E. coli colonies were grown overnight in LB supplemented with the appropriate antibiotic. (2) 1.5 to 3.0 ml of culture was pelleted by centrifugation at 10,000 x G.

Steps 1 to 3 for S. cerevisiae: (1) 3–5 ml of S. cerevisiae transformants (pooled) were harvested from dropout media plates ~5 days after assembly. (2) The cells were pelleted by centrifugation at 3000 x G for 10 min. (3) The P1 resuspension buffer consisted of 240 µl P1 with 5 µl of 1.4 M b-Mercaptoethanol and 5 µl Zymolyase solution (Zymolyase solution: 200 mg Zymolyase 20T (BioShop, catalog number: ZYM001.100), 9 ml H2O, 1 ml 1 M Tris pH 7.5, 10 ml 50% glycerol, stored at -20°C). Following resuspension, the cells were incubated at 37°C for 60 min ahead of the P2 step.

Steps 1 to 3 for P. tricornutum: (1) P. tricornutum transformants were repatched at least twice and grown to high density on ¼-salt L1 plates supplemented with the appropriate antibiotic(s) (i.e., 100 µg/ml nourseothricin or zeocin for single selection, double selection plates contained both antibiotics at 50 µg/ml). On the second passage, colonies were patched as large streaks to ensure there were enough cells for DNA isolation – typically, a single plate would be used to passage 6 colonies that were struck out to each cover approximately a sixth of the plate. Plates were grown until a reasonable cell density was achieved, which typically took 4 days for nourseothricin plates and upwards of 6 days for zeocin, 2-FA, or double-selection plates. (2) Cells were scraped and resuspend in 250 µl of resuspension buffer, which consisted of 245 µl of P1 (Qiagen, catalog number: 19051) and 5 µl of alcalase (i.e., 15 mAnson, Sigma-Aldrich, catalog number: 126741). (3) The resuspended cell mixture was vortexed for 1 to 3 seconds, then placed in a heating block set to 56°C for 10 min – this is within the optimal temperature range for alcalase activity, according to the manufacturer. After 10 min passed, the cells were vortexed again for 1 to 3 seconds to ensure there were no cell clumps.

For E. coli-derived episomes smaller than 11 kb, we used column purification kits to isolate and purify DNA (New England Biolabs, catalog number: T1110). DNA was eluted with ddH2O equilibrated to 56°C.

To isolate a high concentration of pSC5 (~56 kb) from E. coli, alkaline lysis was performed using 27 ml of culture (1.5 ml culture per lysis reaction, 18 reactions in total). The DNA pellets were resuspended with ~25 µl of sddH2O and combined into a single 1.5 ml tube (~460 µl total). Half of the isolated DNA was then digested using SacI-HF, as described below. Then, a tenth volume of sodium acetate (3 M, pH 5.2) and two volumes of ice-cold 100% ethanol were added to the samples. The solutions were inverted to mix and placed into a -80°C freezer for an hour before performing centrifugation at 16,000 x G for 10 min at 4°C. The DNA pellets were then washed twice with 500 µl of ice-cold 70% ethanol before being decanted, dried, and resuspended with 50 µl of sddH2O. This additional sodium precipitation step was performed to further concentrate the samples and reduce the carryover of contaminants, particularly from the digest reaction.

DNA concentrations were measured by fluorometry using the DeNovix dsDNA Broad Range Assay (FroggaBio, catalog number: DSDNA-BROAD-EVAL) and purity ratios were assessed using a Nanodrop spectrophotometer. For the experiments shown in Figs. 2 to 4, samples were adjusted to be equimolar prior to conducting electroporation.

Restriction digests of episomes
Circular pPtGE31_ΔPtR was isolated a P. tricornutum transformant and recovered in EPI300 E. coli. After isolating the episome from E. coli, approximately 10 µg was digested using 3 µl of EcoRI-HF (New England Biolabs, catalog number: R3101) at 37°C for 1 hour. An additional reaction was conducted where 10 µg was digested using 3 µl of EcoRI-HF and 3 µl of mung bean nuclease (New England Biolabs, catalog number: M0250) at 37°C for 1 hour. Complete digestion was confirmed by agarose gel analysis, then samples were purified using a DNA Cleanup Kit (New England Biolabs, catalog number: T1030).

Circular pSC5 was isolated from EPI300 E. coli as described above. Prior to conducting the sodium acetate precipitation step, approximately 40 µg of pSC5 was digested with 18 µl of SacI-HF (New England Biolabs, R3156) at 37°C for 1 hour. Complete digestion was confirmed by agarose gel analysis, then the sample was purified by sodium acetate precipitation, as described above.

Agarose gel electrophoresis
DNA from plasmid isolations or PCR screens were assayed using 1–2% (w/v) agarose gels stained with ethidium bromide. Uncropped images for all gels presented in this manuscript can be viewed in the Source Data file.

Preparation of Cas9:sgAPT1 complex
HiFi Cas9 nuclease (catalog number: 1081060) and synthetic sgRNA were purchased from Integrated DNA Technologies (IDT) and prepared according to the manufacturer’s guidelines. Briefly, 2 to 4 µg of Cas9 (62 µM) was combined with sgRNA (100 µM) in a 1:1.2 molar ratio and then incubated at room temperature for 20 min to form the Cas9:sgRNA complex. After incubation, the complex was set on ice for up to 1 hour, during which the P. tricornutum cells were prepared for electroporation. Once the cells were ready, 40 µg of ssssDNA was added to a 50 µl preparation of the cells, followed by the Cas9:sgRNA complex. The mixture was pipetted up-and-down a few times before transferring to an electrocuvette and promptly pulsed. For the co-transformation experiment, we added 500 ng of PCR-amplified pPtGE31_ΔPtR to the cells ahead of electroporation.

Yeast assembly of pPtGE31_ShBle
The episome pPtGE31_ShBle was created by yeast assembly of three overlapping fragments. First, the episome pPtGE31_ΔPtR was amplified as two fragments using primers that would add 40 bp overlaps for the promoter and terminator region of the ShBle cassette and split the episome in the HIS3 region. Then, the ShBle cassette was amplified from the p0521s plasmid (Addgene ID: 62862)6 using primers that would add 40 bp overlaps for the insertion site in pPtGE31_ΔPtR (all primers are listed in Supplementary Data 2). Yeast assembly was conducted as previously described39 using ≥ 100 ng of each fragment in place of a bacterial donor.

Electroporation to P. tricornutum
Prior to attempting this methodology, it is worth noting that there is a great degree of variance between electroporation equipment, and this can drastically impact the efficiency of transformation to P. tricornutum. If others attempt this protocol, it may be necessary to initially attempt electroporation with several different parameters to determine which settings work best with different set-ups. For our set-up (500 V, 50 μF, 400 Ω), an empty electrocuvette generates a time constant of ~32 to 35 milliseconds. A neighboring lab has successfully performed electroporation to P. tricornutum using an older electroporator set to the originally described parameters by Zhang and Hu (500 V, 25 μF, 400 Ω)5.

P. tricornutum cells were grown in liquid media or on agar plates for the experiments conducted in Fig. 1 – all successive experiments used cells derived from agar plates. For liquid cultures, 1 ml of cells adjusted to 1–2 × 108 cells/ml were passaged into 50 ml of L1 media and placed in a growth chamber on a shaker set to ~100 rpm. For agar plate cultures, 250 µl of cells adjusted to 1 × 108 cells were spread onto a ½-salt L1 plate with 1% agar (w/v). Liquid and agar plate cultures were grown for 4 days. After 4 days, cells were scraped from the agar plates using 750 µl of L1 and transferred to a 1.5 ml tube. An additional 750 µl of L1 was used to remove any remaining cells from the plate. Then, 10−1 to 10−2 dilutions of the liquid and agar plate cultures were prepared and used to estimate cell density with a hemocytometer. Liquid and agar plate cultures were adjusted to have a final cell density between 2 and 4 × 108 cells/ml; for the liquid cultures, this requires concentrating the cells via centrifugation and resuspension in a smaller volume. Here, the cells are transferred from the flask to a 50 ml conical tube and pelleted by centrifugation at 2000 x g for 10 min at 18°C. The pellet was resuspended with 375 mM D-sorbitol to a concentration between 2 and 4 × 108 cells/ml and transferred to a 1.5 ml tube. Cells derived from agar plates were pelleted by centrifugation at 750 x g for 4 min; here, we used a microcentrifuge, which could pellet the cells at a lower centrifugal speed. As before, the cells were resuspended using 375 mM D-sorbitol to a concentration between 2 and 4 × 108 cells/ml. The final cell concentration was equalized between the liquid and agar plate-derived cultures for every biological replicate.

To spheroplast, 1 mAnson of alcalase (Sigma-Aldrich, catalog number: 126741) was added for every 1 × 108 cells. At this stage, the cells will have been previously concentrated to a density between 2–4 × 108 cells/ml and already resuspended in 375 mM D-sorbitol. Alcalase was diluted 10-times in sddH2O to reduce the pipetting error associated with small volumes. After the addition of alcalase, cultures were placed on a rocking shaker set to ~15 rpm at room temperature for 20 min, during which spheroplasting occurs. Once 20 minutes had passed, the cells were pelleted by centrifugation at 325 x G for 4 min. The supernatant is decanted, and the cells are gently resuspended with 1 ml of ice-cold 375 mM D-sorbitol using a P1000 pipette. Following this step, all materials (i.e., cells, DNA, D-sorbitol, electrocuvettes) should be kept on ice for the duration of this protocol.

The electroporation protocol we used is adapted from Zhang and Hu5, with adjustments made by Kassa et al.18 and Pampuch (unpublished). First, the spheroplasted cells were pelleted by centrifugation at 325 x G for 4 min. The supernatant is decanted, and the pellet is gently resuspended with 1 ml of ice-cold 375 mM D-sorbitol. This process of centrifugation and resuspension is repeated another 2 times, such that in total, the cells will have been resuspended with 375 mM D-sorbitol five times (this includes centrifugation steps prior to and post spheroplasting). After the fifth wash, the cells are spun at 750 x G for 3 min, and a P200 pipette is used to gently remove any remaining supernatant. The cells are gently resuspended with 100 to 200 µl of 375 mM D-sorbitol such that the total volume of resuspended cells will be between ~150 to 250 µl. Into sterile 1.5 ml tubes, 50 µl of cells are aliquoted along with 4 µl (10 µg/µl) of single-stranded salmon sperm DNA (ssssDNA, Sigma-Aldrich, catalog number: D7656) and the DNA to be transferred. Of note, the ssssDNA should be heated to 95°C for 2 to 5 min and then cooled on ice prior to this to ensure it remains single-stranded. We used between 1 to 1000 ng of an 11 kb episome for most transformations in this paper, with 500 ng of this construct appearing sufficient to saturate the electroporation reaction. Negative controls consisted of only ssssDNA in the cell mixture.

Cells are gently pipetted up and down 2 times before transferring to an ice-cold sterile electrocuvette (Fisher Scientific, catalog number: FB102), ensuring that no air bubbles have been transferred and the cells lie evenly across the bottom. The electrocuvette is pulsed in an electroporator (BioRad, Gene Pulser Xcell System, catalog number: 1652660) set to the following parameters – voltage: 500 V, capacitance: 50 μF, and resistance: 400 Ω. The time constant varied from 30 to 32.5 milliseconds for the experiments conducted in Figs. 1–4. Immediately after pulsing, 1 ml of L1 media is gently added to the electrocuvette without disturbing the cells. The electrocuvettes are left to rest at room temperature for 10 min before fully resuspending the cells and transferring the mixture to 10 ml of L1 in 50 ml Falcon tubes. The lids of the Falcon tubes are left loose for air transfer, and the cultures are moved to a growth chamber to recover for 16 to 24 h.

After recovery, the cultures are pelleted by centrifugation at 2000 x G for 10 min at 18°C. The pellet is resuspended with 500 µl of L1 media, then 50 µl (one-tenth), 125 µl (a quarter), and/or 250 µl (half) of the cells are spread onto ¼-salt L1 plates supplemented with the appropriate antibiotic(s).

PEG transformation of P. tricornutum
The optimized PEG transformation protocol is similar to the methods described by Karas et al.6, with some important modifications. First, 250 µl of wild-type cells adjusted to 1 × 108 cells were spread onto a ½-salt L1 plate with 1% agar (w/v). After 4 to 5 days of growth, cells were scraped from the agar plates using 1.5 µl of L1 and transferred to a 1.5 ml tube. The cells were counted via a hemocytometer and adjusted to a concentration of 2–4 × 108 cells; in our experience, the density of the scraped cells typically falls within this range.

For protoplasting, 1 ml of cells was transferred to a 50 ml Falcon tube containing 9 ml of L1 and 1000 µl of alcalase (i.e., 3 Anson); here, alcalase is used in place of the zymolyase, hemicellulase, and lysozyme cocktail previously used by Karas et al.6. This mixture was then placed on a slow rocking shaker (~15 rpm) and incubated at room temperature for up to an hour. The protoplasting efficiency can be estimated by examining the cells at ≥200x magnification during incubation. The majority of cells (>80%) should have formed protoplasts before proceeding to the next step. In our hands, the protoplasts remain intact during this reaction, likely because the cells are incubated in full-salt L1 medium, which has a high osmolarity. A relatively high volume of alcalase is required compared to the electroporation method, which may reflect reduced enzyme activity in the presence of L1 salts.

After incubation, 40 ml of L1 was gently added to the cells, which were then pelleted by centrifugation at 1500 x G for 5 min at 18°C. The supernatant was carefully decanted, and the cells were gently resuspended with 500 µl of L1 followed by 8 µl of ssssDNA (10 µg/µl). For all further centrifugation steps, the pellet should be resuspended gently to avoid mechanically shearing the protoplasts. Then, 250 µl of protoplasts were combined with episomal DNA (circular or linear, 500 ng to 1 μg) and 1 ml of 20% PEG-8000 (w/v; also contains 10 mM Tris pH 8, 10 mM CaCl2, 2.5 mM MgCl2) equilibrated to 30°C.

The protoplast:DNA:PEG suspension was gently mixed through inversion and then incubated at RT for 30 min. After incubation, the mixture was centrifuged for 7 min at 1500 x g. The supernatant was removed and cells were resuspended with 1 ml of L1, and then transferred into a 50 ml Falcon tube containing an additional 9 ml of L1 (total volume of 10 ml). The cells were then recovered in the growth chamber for ~48 h (i.e., 2 days). After recovery, the cells were centrifuged for 5 min at 1500 x g and 18°C. The supernatant was decanted, and the pellet was resuspended in 500 μl of L1, then 125–250 μl of the cells were plated on ¼× L1 supplemented with nourseothricin (100 μg/ml). Plates were left in the growth chamber for at least two weeks before being pictured.

PEG transformation of T. pseudonana
Approximately 1 × 105 cells/ml of T. pseudonana was inoculated into f/2 medium. After 4 days of growth (end of exponential phase), cells were pelleted by centrifugation at 1500 x g for 10 min and resuspended to a concentration of 3 x 108 cells/ml. The same transformation protocol was followed as described for P. tricornutum; however, the protoplasting step was conducted in f/2 medium with differing amounts of alcalase and incubation lengths (Supplementary Fig. 12a). Approximately 500 ng of the 8.2 kb episome pBIG1 was combined with 40 µg of carrier DNA for transformation. Following PEG transformation, the cells were recovered in 10 ml of f/2 media and recovered for 2 days prior to plating. Post-recovery, the cells were centrifuged for 5 min at 1500 x g. The cells were then resuspended in 500 μl of f/2 media, and 250 μl was plated on full-salt f/2 agarose plates supplemented with nourseothricin (100 μg/ml).

Screening P. tricornutum colonies transformed with two fragments – NHR pathway
Non-overlapping fragments from pPtGE31_ShBle were electroporated both individually and simultaneously into P. tricornutum (Supplementary Fig. 3). Cells that were electroporated with both fragments were plated onto ¼-salt L1 plates supplemented with 100 µg/ml nourseothricin or 100 µg/ml zeocin. Transformants that appeared following electroporation with both fragments were passaged twice – first, onto media supplemented with the same antibiotic as the initial transformation, and then onto media supplemented with the alternative antibiotic (Supplementary Table 4). Twenty transformants, ten derived from each initial transformation plate, that demonstrated resistance to both nourseothricin and zeocin were passaged for a third time onto double-selection media (¼-salt L1 supplemented with 50 µg/ml nourseothricin and zeocin). DNA isolation was performed using cells from this third passage, followed by PCR screening.

Screening P. tricornutum colonies transformed with multiple fragments – HDR pathway
Overlapping fragments were adjusted to a final concentration of 56 nM, which is four times more concentrated than the linear episome that was used as a positive control in these experiments (14 nM). Different combinations of the fragments were electroporated to measure the amount of carry-over for the template plasmid, pPtGE31_ShBle. For the experiments exploring the limits of HDR assembly, the molarity of the insert was ~4-5x greater than the episomal fragments. The CytB-eGFP and eGFP-T2A-NAT constructs were commercially synthesized from IDT and PCR amplified to generate ample DNA ahead of assembly; the sequences of these constructs are available in the Source Data file, along with the primers in Supplementary Data 2.

In both series of experiments, cells were plated onto ¼-salt L1 plates supplemented with 100 µg/ml nourseothricin and grown for at least two weeks before passaging 10 colonies from the assembly plates onto the same type of selection plate. The repatched colonies were grown for one week before passaging again for DNA isolation and/or microscopy.

Recovery of P. tricornutum episomes in E. coli
Episomes were isolated from P. tricornutum transformants as described above. For each electroporation reaction, 2 to 4 μl of P. tricornutum DNA was combined with 25 to 50 μl of homemade electrocompetent EPI300 cells (derived from LGC Biosearch Technologies, catalog number: EC02T110) in a sterile 1.5 ml tube. The mixture was gently pipetted twice and then transferred to a sterile 2 mm electrocuvette, ensuring no air bubbles were transferred. The electrocuvette is pulsed in an electroporator set to the following parameters – voltage: 2.5 kV, capacitance: 25 μF, and resistance: 200 Ω. This generated a time constant of 4.9 to 5.2 milliseconds. Electroporated cells were resuspended by pipetting 1000 μl of SOC media up-and-down in the electrocuvette, and then transferred to sterile 1.5 ml tubes to recover at 37°C and 225 rpm for approximately 1 hour. After recovery, half of the reaction volume (~400 μl) was spread across a 1.5% agar (w/v) LB plate supplemented with 15 μg/ml chloramphenicol (pPtGE31_ ΔPtR and pPtGE31_ShBle recovery) or 40 μg/ml gentamycin (pSC5 recovery), which were then transferred to a 37°C incubator. Transformants appeared across the plates within 24 h, with there being a range from 4 to ~100 colonies per transformation.

For recovery of pPtGE31_ΔPtR and pPtGE31_ShBle episomes. Single E. coli colonies were picked and inoculated into 5 ml of LB media supplemented with 15 μg/ml chloramphenicol. Cultures were grown overnight at 37°C and 225 rpm. The following day, 500 μl of culture was inoculated into 4.5 ml of LB media supplemented with 15 μg/ml chloramphenicol and 100 μg/ml L-arabinose. These episomes contain a pCC1BAC backbone, which can be induced to high-copy number replication in EPI300 by adding L-arabinose to the media. Induced cultures were grown for at least five hours prior to episome isolation, with 1.5 ml of culture being used for DNA isolation, as described above.

For recovery of the conjugative plasmid pSC5. DNA isolated from eight P. tricornutum pSC5 transformants were electroporated into EPI300 cells. Four colonies per transformation were inoculated into 5 ml of LB supplemented with 40 μg/ml gentamycin. Additionally, EPI300 strains carrying pSC5 (positive control) or pSAP (recipient cell line) were inoculated from glycerol stocks into 5 ml of LB supplemented with 40 μg/ml gentamycin (pSC5) or 15 μg/ml chloramphenicol (pSAP). The cultures were placed at 37°C and 225 rpm. After the cultures reached saturation (~18 h), 125 μl of the pooled E. coli transformants and pSC5 cultures were inoculated into 5 ml of LB supplemented with 40 μg/ml gentamycin. For the pSAP culture, 5 ml of culture was diluted into 45 ml of LB supplemented with 15 μg/ml chloramphenicol. The cultures were grown at 37°C and 225 rpm for 2 h. Then, the cultures were pelleted by centrifugation at 3000 x G for 12 min at 10°C, decanting the supernatant post centrifugation. The pooled transformants and pSC5 cultures were each resuspended with 100 μl of 10% glycerol, whereas the 50 ml pSAP culture was resuspended with 1000 μl of glycerol. Strains were kept on ice during these steps to cease bacterial growth.

For conjugation, 100 μl of the donor cells (transformant pools [x8] and pSC5) were mixed with 100 μl of the recipient cells (pSAP). The cells were mixed by pipetting up and down 4 times before transferring the whole mixture to a 2% agar (w/v) LB plate. Additionally, 100 μl of the recipient was plated by itself to serve as a negative control. Conjugation plates were incubated at 37°C for 1 hour. Then, plates were scraped using 2 ml of sddH2O. The scraped cells were vortexed for 5 seconds to ensure any clumps were dispersed. Transconjugants were spot plated onto 1.5% agar (w/v) LB plates supplemented with 40 μg/ml gentamycin and 15 μg/ml chloramphenicol with dilutions ranging from 100 to 10-8.

Sequencing and alignment
Amplicons generated by amplifying the episome termini region in P. tricornutum transformants were sent for Sanger sequencing (London Regional Genomics Centre). Episomes isolated from E. coli were sent for Oxford Nanopore whole-plasmid sequencing (Plasmidsaurus Inc. and Flow Genomics). The resulting sequences were manually aligned to the respective template episomes. Regions that could not be manually aligned were queried using a local alignment algorithm (EMBOSS water tool)40 to search for homology in the episome. Sequences that could not be aligned manually or algorithmically were queried using BLASTn (NCBI)41.

Analyses of P. tricornutum transformation efficiency
The total number of colonies per transformation reaction was estimated by manually counting the number of CFUs per plate and then multiplying this value by 2 or 10, depending on whether half or a tenth of the total reaction was plated, respectively. In instances where the transformation plate was too dense to reliably count by eye, five 1 cm2 quadrants were sampled from the plate and counted using a dissecting microscope with 10x magnification. Here, the total number of CFUs was estimated by averaging the number of colonies per quadrant and multiplying this value (i.e., colonies per cm2) by the total surface area of the petri dish.

Transformation efficiencies were estimated by dividing the total number of CFUs by the number of cells per reaction. When graphically comparing transformation reactions, all calculations and plotting were done in RStudio (version 2025.05.1+513); otherwise, we used an Excel spreadsheet (version 16.103.4) to keep track of data and tabulate transformation efficiencies (available in the Source Data file).

Statistics and reproducibility
For experiments in which transformation efficiency was quantitatively assessed (Fig. 1e, f), four to five independent biological replicates were performed to account for variability between electroporation experiments and to enable statistical analysis. For all other experiments, which were designed as qualitative proof-of-concept assays (e.g., assessment of NHR and HDR assembly), at least two independent biological replicates were performed. Where appropriate, representative plate images from a single biological replicate are shown.

Differences in transformation efficiency across the four treatment groups (plate-derived untreated, plate-derived alcalase-treated, liquid-derived untreated, and liquid-derived alcalase-treated) were statistically assessed using a two-sided Kruskal-Wallis test. Pairwise comparisons between selected groups (LU vs. PS, LS vs. PS, PU vs. PS) were performed using Wilcoxon rank-sum tests with Bonferroni correction for multiple comparisons.

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "This manuscript presents a significant advancement in genetic engineering tools for diatoms, focusing on optimized protocols for DNA and ribonucleoprotein (RNP) complex delivery to Phaeodactylum tricornutum. The authors demonstrate that spheroplasting via alcalase treatment greatly enhances transformation efficiency for electroporation and PEG-mediated delivery. The study introduces the concept of Diatom In Vivo Assembly (DIVA), showing that episomes, including those up to 55.6 kb, can be assembled directly within diatom cells through non-homologous end joining (NHEJ) or homologous recombination (HR). Additionally, they present an electroporation-based protocol for DNA-free genome editing using Cas9 RNPs, achieving successful gene knockouts and even co-delivery of RNPs with episomes. These developments are positioned to support synthetic biology efforts, such as the Pt-syn 1.0 project, and can be applied to other diatom species, including Thalassiosira pseudonana. The work is clearly described, the data are compelling, and the implications for synthetic biology are profound. The following points can be considered to enhance the clarity of the paper:\nPage 3, Lines 103–104:\nData confirming the absence of genomic integration and demonstrating that the episome is maintained in a circular form should be provided to support these sentences. “Data now shown” should be avoided.\nPage 4, Line 116–:\nIn order to substantiate the hypothesis that diatom cell morphology and cell wall integrity affect transformation efficiency via electroporation, experiments involving alcalase treatment should be conducted on oval, fusiform, and triradiate cells. While the following section presents data from liquid cultures, which are likely to contain predominantly fusiform cells, this should be discussed in the former section for consistency and clarity.\nFurthermore, more detailed microscopic images showing the morphological changes induced by alcalase treatment would improve understanding of this process. What morphological changes were observed in fusiform and triradiate cells upon treatment? Were those the reasons for lower transformation efficiency?\nFigure 2e:\nPlease comment on the absence of a band at around 486 bp in lanes 3, 4, and 5 in the text. Could the integration of the introduced DNA fragments into the genome result in resistance to the selection antibiotics?\nPage 16, Line 501–:\nTo ensure the purpose of this experiment is immediately clear to the readers, it would be helpful to briefly state the purpose of the co-delivery of RNPs and episomes in this paragraph.\nFigure 7:\nAlthough Figure 7 suggests the potential for the targeted insertion of foreign DNA at arbitrary loci in the diatom genome, this possibility has not been demonstrated experimentally in the current study. Although this is a plausible and exciting prospect, the figure should be omitted to avoid misleading readers.", "reviewer_2": "General Comments:\nThis manuscript presents promising improvements in DNA delivery protocols for P. tricornutum, particularly through the use of spheroplasting in combination with electroporation and polyethylene glycol (PEG)-mediated transformation. The authors demonstrate improved transformation efficiency using both circular and linearized plasmids, and successfully apply ribonucleoprotein (RNP) complexes of Cas9 and gRNA for gene editing. Notably, they provide the first, exciting evidence of in vivo DNA fragment assembly in diatoms.\nHowever, in its current form, the manuscript primarily presents an optimization of existing transformation protocols. While the authors demonstrate successful delivery of various constructs into diatom cells, they do not investigate the fate of the introduced DNA within the algal cells—specifically, whether integration into the genome occurs, the copy number and stability of episomal plasmids, or the potential for epigenetic silencing. Additionally, the novel and potentially impactful observation of in vivo DNA assembly in algae is acknowledged but not explored in depth.\nAs such, the manuscript may be more appropriate for a technically oriented journal. Alternatively, it could be significantly strengthened by expanding on the mechanisms and biological implications of in vivo DNA assembly in diatoms.\nBelow are specific comments and suggestions for revision:\nMajor comments:\nOverstatements\nThe authors claim that DNA delivery is the major challenge in diatom transformation. While this is true for several diatom species, it does not hold for Phaeodactylum tricornutum, which can be efficiently transformed using multiple established methods. In contrast, a more significant challenge in P. tricornutum is the frequent epigenetic silencing of transgenes. Furthermore, P. tricornutum lacks a silica cell wall, making it particularly well-suited for spheroplasting-based approaches.\nThe inclusion of Thalassiosira pseudonana is also unconvincing. As the other main model diatom species, T. pseudonana has a well-documented history of successful transformations, and it is unclear how the current protocol represents a major advancement for other species.\nThe authors further propose that de novo diatom in vivo assembly (DIVA) could replace traditional cloning approaches in E. coli and S. cerevisiae, which they describe as time-consuming. However, this claim is not well substantiated. The data presented show that in vivo assembly in diatoms is prone to errors, in contrast to the reliable transformation with pre-assembled plasmids. Moreover, assembled episomes still require purification and transformation into E. coli for verification. This statement also does not account for the lengthy selection process in diatoms, which typically takes around three weeks, compared to the single-day turnaround in E. coli. Instead of positioning DIVA as a general replacement for bacterial or yeast cloning, we suggest that the authors propose a more practical and specific application where DIVA could offer a clear advantage.\nCo-transformation with RNP Cas9-gRNA complexes and episomal resistance marker: The manuscript presents this strategy as a solution for targeting genomic loci that lack a selectable phenotype. However, with a reported co-transformation efficiency of only 10%, the majority of resulting colonies (~90%) are false positives. The authors might consider exploring an alternative strategy—such as co-delivery of multiple RNP complexes targeting different loci—similar to the approach described in Serif et al., 2018.\nExamples of overstatements:\nLine 28: The sentence overgeneralizes the adaptability of the protocol. Thalassiosira pseudonana is also a model diatom species with a small genome, rapid growth, and generally accesible for genetic transformation. Given the differences between species and that electroporation was only tested in P. tricornutum (and most effective on plates), caution should be used when implying broad adaptability.\nLine 64: “Conjugation requires the creation and maintenance of a donor strain of Escherichia coli, which must contain a suitable broad-host-range conjugative plasmid in addition to the episome of interest.” While this statement is technically accurate, it is unclear why the authors present it as a drawback. The creation and maintenance of donor E. coli strains is a standard molecular biology procedure and typically not considered a major limitation.\nLine 89: Reiteration of line 28 comment: the claim that these methods can be broadly applied to other diatoms should be more cautious and supported with data or omitted.\nGeneral note: The data suggest that spheroplasting improves transformation mainly on plates, with limited benefit in liquid culture. This is important because many diatom species cannot grow on solid media.\nUneven quality across the manuscript:\n- The final two sections—PEG transformation and Thalassiosira pseudonana transformation—are notably underdeveloped compared to the rest of the experimental work. These sections appear rushed, lacking sufficient detail, proper quantification, and clear presentation of results. To maintain consistency and clarity, the authors should either expand and revise these sections to match the depth and quality of the earlier parts of the manuscript or consider omitting them altogether.\nOpen questions:\nWould the addition of telomeric sequences help stabilize the linearized plasmids and prevent their circularization in diatom cells?\nCould the authors clarify why adenine supplementation is necessary following PtAPT knockout? What is the underlying metabolic or genetic rationale?\nAdditional line-by-line comments:\nLine 26: The term “PEG” is used in the Abstract without being defined. Please define “polyethylene glycol” at first mention.\nLine 63: The transformation rate is incorrectly written as “4 transformants per 10⁻⁴ cells”. It should read “4 transformants per 10⁴ cells” for consistency with line 58.\nLine 69: More background and context on PEG-mediated transformation in diatoms is needed. Currently, the discussion is too brief for readers unfamiliar with the technique.\nLine 70: Correct “≤ 1 transformant per 10⁻⁸ cells” to “≤ 1 transformant per 10⁸ cells”.\nLine 87: The manuscript inconsistently uses the terms spheroplasting and protoplasting. These are sometimes used interchangeably and sometimes as distinct processes. Given the title refers specifically to “spheroblasted cells,” a clear and consistent definition is essential.\nLine 97: The authors note that a neighbouring lab used older electroporation parameters and that the switch from 25 to 50 µF may relate to the equipment used. This raises the question of whether the improved transformation efficiency is protocol-specific or simply equipment-dependent. Please clarify.\nLine 101: Plasmid pPtGE31_ΔPtR with ID 236260 is not found on Addgene. Please upload the plasmid map and sequence before any resubmission.\nLine 176: The use of 500 ng of a PCR-amplified 11-kb construct should be mentioned earlier in the text, especially when referring to Figure 1.\nLine 256: The claim that linear pSC5 is three times more efficient than circular is not clearly supported by Figure 3B, where colony counts appear similar (5 vs. 6 CFUs). Please clarify.\nLine 302: Plasmid pPtGE31_ShBle with ID 236261 is not found on Addgene. Please upload the plasmid map and sequence before any resubmission.\nLine 393: There is no Figure S6C. This likely refers to Figure S7C—please correct.\nLine 539: A brief conclusion summarizing the main findings and potential applications would strengthen the end of the manuscript.\nLine 550: The difference in DNA quantities used (500 ng linear vs. 25 µg circular) is substantial. Please explain how these values were chosen.\nLine 554: Early in the manuscript, alcalase dosage is reported in mAnson units per 10⁸ cells, but here it is given in µl. Please standardize units throughout.\nLine 558: The text states that 40 µg of carrier DNA improved electroporation efficiency ~15-fold, but no results are shown for PEG transformation. Was this amount used in PEG protocols? Was a similar improvement observed?\nLine 565: The final optimized PEG protocol is not clearly presented. Please summarize the final conditions used and their effectiveness.\nLine 566: There is no Figure S2D, despite the reference. Please correct or include the figure.\nLines 565–571: This paragraph lacks quantitative data and clarity. It is difficult to assess the advantage of PEG transformation without specific efficiency data.\nLine 574: Reiterate the need to clearly outline the final optimized conditions.\nLine 577: Please include the size of the episome used here, as done in the Materials and Methods section.\nLines 578–581: The results described are vague. Clarify what “some degree of growth” means and how harsher protoplasting conditions influenced colony formation.\nLine 588: The phrase “a higher proportion of protoplasted cells are visible…” is vague. Provide more concrete observations or quantification, if available.\nLine 593: There is no discussion on partial vs. complete cell wall removal. Is this dependent on enzyme concentration? Do cells survive full wall removal?\nLine 611: It is true that plated cells yielded the highest transformation rates, but based on Table S2, efficiencies are mostly in the 10⁶–10⁷ range, with only treated plated cells showing higher values (1.19 x 10⁴). The benefit of alcalase treatment seems to apply mainly to plated cells.\nLine 612: The manuscript states that transformation is “efficient,” but no numerical threshold is provided. Please define what constitutes efficiency in this context.\nLine 645: “Several-fold more efficient” should be quantified.\nLine 701: Only PEG transformation was adapted to T. pseudonana. The claim that these methods hold great promise for other diatoms lacks sufficient supporting evidence.\nLine 866: The note about keeping materials on ice is useful and should be moved earlier in the protocol section for clarity.\nLine 880: The DNA input range (“1 to 1000 ng”) is very broad. Consider adding a note that for ~11-kb episomes, 500 ng appears to saturate the reaction.\nLine 904: This line again references equipment differences affecting electroporation settings. Consider addressing this earlier in the Methods for better context.\nFigure Comments:\nFigure 1b: Why is the image shown with 100 mAnson if the best treatment was achieved with 1 mAnson? This is confusing and needs explanation.\nFigure 1e and f: Statistical analysis should be included. Please clarify what alcalase treatment was used and provide full experimental conditions (e.g., DNA concentration, treatment duration) so that the figure is self-contained.\nFigure 1e: As noted above, experimental details (e.g., DNA concentration, alcalase protocol) are missing. These are essential for interpretation.\nFigure 1f: Clarify whether this experiment was done in liquid culture or on plates, and specify the alcalase treatment used.\nFigure 2c–e: Please clarify the annotations in panels d and e to clearly indicate which PCR primer pairs were used, with explicit reference to panel c.\nFigure 3e is not clearly explained. Presumably, it should refer to line 269: “conjugation was successful for seven out of eight E. coli pools”. If it is the case, then labelling C1-C4 in both cases is confusing.\nFigure 5: Part a shows three schemes of possible assembly. Part b does not correspond to part a. Please adjust for clarity.", "reviewer_3": "I co-reviewed this manuscript with one of the reviewers who provided the listed reports. This is part of the Nature Communications initiative to facilitate training in peer review and to provide appropriate recognition for Early Career Researchers who co-review manuscripts."}, "author_response": "Our responses are in purple.\nIn addition to the comments provided by Reviewers 1 to 3, we also had a colleague, Dr.\nDavid Edgell, read our manuscript. We have made additional changes in accordance with\nDr. Edgell’s suggestions; notably, it was not correct to refer to the assembly methods as\nnon-homologous end joining and homologous recombination. A better term for these\nprocesses is non-homologous repair and homology-directed repair, since we do not know\nthe exact underlying DNA repair process governing episomal assembly in P. tricornutum.\nWe also caught some additional small errors upon carefully assessing the manuscript.\nReviewer #1 (Remarks to the Author):\nThis manuscript presents a significant advancement in genetic engineering tools for\ndiatoms, focusing on optimized protocols for DNA and ribonucleoprotein (RNP) complex\ndelivery to Phaeodactylum tricornutum. The authors demonstrate that spheroplasting via\nalcalase treatment greatly enhances transformation efficiency for electroporation and\nPEG-mediated delivery. The study introduces the concept of Diatom In Vivo Assembly\n(DIVA), showing that episomes, including those up to 55.6 kb, can be assembled directly\nwithin diatom cells through non-homologous end joining (NHEJ) or homologous\nrecombination (HR). Additionally, they present an electroporation-based protocol for DNA-\nfree genome editing using Cas9 RNPs, achieving successful gene knockouts and even co-\ndelivery of RNPs with episomes. These developments are positioned to support synthetic\nbiology efforts, such as the Pt-syn 1.0 project, and can be applied to other diatom species,\nincluding Thalassiosira pseudonana. The work is clearly described, the data are\ncompelling, and the implications for synthetic biology are profound. The following points\ncan be considered to enhance the clarity of the paper:\nWe sincerely thank Reviewer 1 for their positive and encouraging feedback. We are\ngrateful that they found the manuscript to be of interest and appreciate their constructive\nfeedback for enhancing clarity. We have carefully considered each point raised and have\nrevised the text accordingly. Below, we provide a detailed response to each comment.\nPage 3, Lines 103–104:\nData confirming the absence of genomic integration and demonstrating that the episome is\nmaintained in a circular form should be provided to support these sentences. “Data now\nshown” should be avoided.\nThese are valid points; we should have provided more data to support these statements.\nTo address the first point (i.e., genomic integration), we have performed a\nplasmid/episome loss experiment by serially passaging four pPtGE31_ ΔPtR\ntransformants in non-selective media. The transformants chosen correspond to algal\ncolonies 1 and 2 in Fig. 2D and 1 and 8 in Fig. 2E. After seven passages in liquid non-\nselective media, serial dilutions of the cultures were then plated onto solid non-selective\nmedia to obtain single colonies. For each transformant, 100 single colonies were then re-\npatched onto plates supplemented with nourseothricin. If genomic integration of the\nepisome had occurred, we would expect a high proportion of the single colonies to retain\nresistance. Conversely, if the episomes are maintained extrachromosomally, they should\nbe lost over successive generations without selective pressure, and thus only a low\nproportion of colonies would retain resistance. As expected, only a small fraction of\n1\nscreened colonies retained resistance (Supplementary Table S4), demonstrating that\ngenomic integration had not occurred in the assayed transformants.\nWith this in mind, we understand that genomic integration can still occur when\ntransforming episomes, regardless of the delivery method (e.g., conjugation or\nelectroporation); however, this outcome is probabilistically less likely. For integration to\nsuccessfully yield a nourseothricin-resistant transformant, the construct must not only\nintegrate into a DSB in the genome, but the region of integration must also be\ntranscriptionally active. Comparatively, the episome just needs to make it to the nucleus to\nyield a successful transformant (generally speaking). This is illustrated in Supplementary\nFig. S1, where we transformed equimolar amounts of the episome pPtGE31_ΔPtR (~11\nkb) and the nourseothricin resistance cassette (~1.4 kb) into P. tricornutum cells. Our work\nwith pSC5 (56 kb) pPtGE31_ΔPtR demonstrated how construct size can significantly\nimpact the efficiency of electroporation. Despite the NAT cassette being nearly tenfold\nsmaller than pPtGE31_ΔPtR, it produced over tenfold fewer colonies, highlighting the\nlower likelihood of successful genomic integration compared to episomal delivery.\nWe have clarified this point in the revised manuscript by adding the following section:\n“Evaluation of genomic integration\nWe have performed a plasmid/episome loss experiment by serially passaging four\npPtGE31_ ΔPtR transformants in non-selective media to check for genomic integration.\nThe transformants chosen correspond to algal colonies 1 and 2 in Fig. 2D and 1 and 8 in\nFig. 2E. After seven passages in liquid non-selective media, serial dilutions of the cultures\nwere then plated onto solid non-selective media to obtain single colonies. For each\ntransformant, 100 single colonies were then re-patched onto plates supplemented with\nnourseothricin. If genomic integration of the episome had occurred, we would expect a\nhigh proportion of the single colonies to retain resistance. Conversely, if the episomes are\nmaintained extra chromosomally, they should be lost over successive generations without\nselective pressure, and thus only a low proportion of colonies would retain resistance. As\nexpected, only 2-7% of screened colonies retained resistance (Supplemental Table S4),\ndemonstrating that genomic integration had not occurred in most assayed transformants.”\nTo address the second point (i.e., plasmid recircularization), we have added additional\ndata (Supplementary Fig. S2) for clarity. The EPI300 E. coli strain cannot propagate non-\ncircular DNA. Therefore, if the algal episomes were maintained linearly, they could not be\nrecovered in EPI300. All of our episomal analyses (e.g., sequencing, conjugation assays)\ninvolved recovery of the DNA in EPI300, but to further illustrate this point, we\nelectroporated PCR-amplified pPtGE31_ΔPtR that had been DpnI treated (to get rid of any\nremnant circular template) into EPI300. This yielded zero colonies, as expected, whereas\nelectroporation of pPtGE31_ΔPtR recovered from algal transformants (which have low\ncopy/poor DNA yield comparatively) yields hundreds of colonies.\nIn addition, we modified text in result section to address this to read as follows:\n“Our initial attempts at electroporation using a previously described protocol16 that was\nadapted from the methods of Zhang and Hu5 yielded no transformants. After several failed\nattempts, we discovered that adjusting the electroporator capacitance from 25 to 50 µF\nyielded transformants for both episomes and marker cassettes, though efficiency was\nhighly variable between experiments (Supplementary Table S1, Exp. 1 to 81). We used\nthis method to deliver a linear 11 kb PCR-amplified episome, pPtGE31_ΔPtR (Addgene\nlinear episome, or be converted into a circular episome within the P. tricornutum nucleus.\nWe were able to recover DNA from these algal transformants in EPI300 E. coli, indicating\nthat the linear episome had been circularized in the algal nucleus following electroporation\n(Supplementary Fig. S2). Whole-plasmid sequencing of four E. coli colonies confirmed that\ncircularization had occurred via fusion of the construct’s termini.”\nPage 4, Line 116–:\nIn order to substantiate the hypothesis that diatom cell morphology and cell wall integrity\naffect transformation efficiency via electroporation, experiments involving alcalase\ntreatment should be conducted on oval, fusiform, and triradiate cells. While the following\nsection presents data from liquid cultures, which are likely to contain predominantly\nfusiform cells, this should be discussed in the former section for consistency and clarity.\nWe thank the reviewer for this insightful suggestion. Unfortunately, we cannot readily\ncontrol cell morphology in P. tricornutum, and triradiate cells are particularly rare in our\nstrain, making systematic experiments with all morphotypes impractical. We have\npreviously attempted to propagate cultures from single colonies with a particular\nmorphology before, but found that single colonies will differentiate/switch morphotypes\nrapidly. Nevertheless, our microscopy observations confirm that all cell types can be fully\nprotoplasted with alcalase (can see remnant frustules that appear following the alcalase\ntreatment). We have included additional microscopy images to further illustrate these\nobservations (in accordance with your suggestion below; Supplementary Fig. S3).\nWe have revised text in the results section to clarify this point to read as follows:\n“When this electroporation protocol was repeated using cells that had reached the late\nstationary phase of growth, we obtained up to 272 transformants per 108 cells with the\nsame 11 kb PCR-amplified episome (Supplementary Table S1, Exp. 82 to 89). This\nrepresented a nearly 20-fold increase in efficiency when compared to transformations with\nearly-log phase cultures (Exp. 80 and 81). Stationary phase cultures demonstrated\nnoticeable differences in cell phenotype when compared to early-log phase cultures, with\nthere being a higher abundance protoplasts (i.e., diatoms lacking a cell wall) in the older\ncultures. P. tricornutum cells can appear in three different morphotypes – fusiform, oval,\nand triradiate – and can switch morphotypes in response to environmental stimuli17,18. All\nmorphotypes can generate protoplasts if the cell wall is sufficiently weakened. Ultimately,\nthis observation led us to hypothesize that cell morphotype and cell wall integrity may\ninfluence electroporation transformation efficiency.\nCell morphotype cannot be readily controlled in P. tricornutum cultures as this species\ndisplays an immense amount of phenotypic plasticity; however, agar-plated cultures of\nCCAP 1055/1 tend to have a higher proportion of oval-type cells (~20-30% oval, ~70-80%\nfusiform) compared to liquid cultures (<10% oval, >90% fusiform). Thus, to test this\nhypothesis, we spheroplasted early-log phase P. tricornutum cells harvested from agar\nplates. Plate-derived cells were spheroplasted using the enzyme alcalase, a serine\nprotease that has been shown to degrade the proteinaceous components of the P.\ntricornutum cell wall19. We observed that 100 µl of alcalase at an activity of approximately\nderived cells when resuspended in 375 mM D-sorbitol (Fig. 1A and B). The protoplasted\ncells were very fragile and prone to rupture from osmotic pressure or physical damage;\ntreatment of 3 x 108 cells with 10 µl of alcalase (30 mAnson units) was enough to\n3\nprotoplast some cells, whilst leaving others with partially degraded cell walls (i.e.,\nspheroplasts).”\nFurthermore, more detailed microscopic images showing the morphological changes\ninduced by alcalase treatment would improve understanding of this process. What\nmorphological changes were observed in fusiform and triradiate cells upon treatment?\nWere those the reasons for lower transformation efficiency?\nThis is an excellent suggestion, we have added text and microscopy images for cells\ntreated with 0.1, 1, and 10 mAnson alcalase per 108 cells (corresponds to exp. in Fig. 1C)\nto demonstrate the morphological changes induced by the various alcalase treatments\n(Supplementary Fig. S3). We very rarely see triradiate forms in healthy wild-type cultures\nso we couldn’t capture images of this particular morphotype (though we have seen\ntriradiate frustules following spheroplasting treatments), but both the fusiform and oval\nmorphotypes can be visibly protoplasted by alcalase. This is best evidenced by the\nappearance of fusiform and oval ‘shells’ (i.e., empty frustules) in the 1 and 10 mAnson\ntreated cells – these ‘shells’ become increasingly more visible as the strength of the\nalcalase treatment increases. We believe the lower transformation efficiency with liquid\ngrown cultures has more to do with other phenotypic changes that accompany growth on\nsolid media. It’s worth noting that a similar phenomenon occurs during conjugation;\nplated P. tricornutum cells are more efficient than liquid cultured cells (Karas et al, 2015;\ndoi: 10.1038/ncomms7925).\nFigure 2e:\nPlease comment on the absence of a band at around 486 bp in lanes 3, 4, and 5 in the\ntext. Could the integration of the introduced DNA fragments into the genome result in\nresistance to the selection antibiotics?\nWe thank the reviewer for this insightful question. The absence of the 486 bp band in lanes\n3, 4, and 5 is most likely due to DNA degradation or deletions occurring at the episomal\ntermini, either before (e.g., exonuclease activity) or during the non-homologous repair\nprocess. While integration of the introduced DNA fragments into the genome could in\nprinciple confer resistance, the presence of the other amplicon suggests that the construct\nis being maintained episomally. This amplicon is located in the CEN/ARS region, which is\n~3 kb away from the other amplicon (i.e., the site of termini fusion) and ~7.5 kb away from\nthe nat cassette in the linear construct. We have added text to this section to address this.\nPage 16, Line 501–:\nTo ensure the purpose of this experiment is immediately clear to the readers, it would be\nhelpful to briefly state the purpose of the co-delivery of RNPs and episomes in this\nparagraph.\nThis is an excellent suggestion, we have added text to the revised manuscript to clearly\nstate the purpose of these co-delivery experiments. Specifically we added this sentence:\n“The aim of this experiment was to develop protocol that could significantly improve\nscreening for gene deletions that do not produce an obvious phenotype.”\nFigure 7:\nAlthough Figure 7 suggests the potential for the targeted insertion of foreign DNA at\narbitrary loci in the diatom genome, this possibility has not been demonstrated\n4\nexperimentally in the current study. Although this is a plausible and exciting prospect, the\nfigure should be omitted to avoid misleading readers.\nWe agree with this statement; the inclusion of Fig. 7 is too pre-emptive for this manuscript.\nThe figure has been removed and the text has been revised accordingly.\nWe thank the Reviewer #1 again for their thoughtful comments and for helping us improve\nthe manuscript.\nReviewer #2 (Remarks to the Author):\nGeneral Comments:\nThis manuscript presents promising improvements in DNA delivery protocols for P.\ntricornutum, particularly through the use of spheroplasting in combination with\nelectroporation and polyethylene glycol (PEG)-mediated transformation. The authors\ndemonstrate improved transformation efficiency using both circular and linearized\nplasmids, and successfully apply ribonucleoprotein (RNP) complexes of Cas9 and gRNA\nfor gene editing. Notably, they provide the first, exciting evidence of in vivo DNA fragment\nassembly in diatoms.\nHowever, in its current form, the manuscript primarily presents an optimization of existing\ntransformation protocols. While the authors demonstrate successful delivery of various\nconstructs into diatom cells, they do not investigate the fate of the introduced DNA within\nthe algal cells—specifically, whether integration into the genome occurs, the copy number\nand stability of episomal plasmids, or the potential for epigenetic silencing. Additionally, the\nnovel and potentially impactful observation of in vivo DNA assembly in algae is\nacknowledged but not explored in depth.\nAs such, the manuscript may be more appropriate for a technically oriented journal.\nAlternatively, it could be significantly strengthened by expanding on the mechanisms and\nbiological implications of in vivo DNA assembly in diatoms.\nWe sincerely thank Reviewers #2 and #3 for their combined extensive feedback, it has\nhelped strengthen and refine our work. We will provide more detailed responses below to\nthe specific comments and suggestions, but first, we wanted to address some of the\ncomments above, starting with:\n“While the authors demonstrate successful delivery of various constructs into diatom cells,\nthey do not investigate the fate of the introduced DNA within the algal cells—specifically,\nwhether integration into the genome occurs, the copy number and stability of episomal\nplasmids, or the potential for epigenetic silencing.”\nThis is a valid and important criticism, and we have taken steps to address each aspect as\nfollows:\nGenomic integration: We have added additional section in the revised manuscript to\naddress whether genomic integration occurs (see also Reviewer #1’s comments). In brief,\nwhile genomic integration of the episome can occur, it is probabilistically far less likely than\nepisomal maintenance. Successful integration would require multiple chance events: the\nepisome must first reach the nucleus, and then must integrate into the genome at a\nsporadic double-strand break (DSB) site located coincidentally within a transcriptionally\nactive genomic region; conversely, for episomal maintenance, the episome just has to\n5\nreach the nucleus and circularize, as our work has revealed, for successful transformation.\nAn example of this can be seen in Supplementary Fig. S1, where we transformed\nequimolar amounts of the episome pPtGE31_ΔPtR (~11 kb) and the nourseothricin\nresistance cassette (~1.4 kb) into P. tricornutum. Despite being nearly tenfold smaller (=\nhigher transformation efficiency), the cassette yielded over tenfold fewer colonies than the\nepisome, underscoring the lower likelihood of successful genomic integration due to the\nneed for a DSB in a transcriptionally active region.\nCopy number and stability: We did not explicitly investigate episomal copy number or\nstability following electroporation or PEG-mediated transformation, as we do not anticipate\nsignificant differences compared with other DNA delivery methods – particularly in\ncomparison to biolistics, which also relies on physical force to introduce DNA into cells.\nThe episomes used in this work have been previously characterized and shown to be\nstable (Slattery et al., 2018; doi: 10.1021/acssynbio.7b00191), and past work has also\nshown that episomal DNA in P. tricornutum is maintained at low copy number within the\nnucleus (Karas et al., 2015; doi: 10.1038/ncomms7925). We did report on the existence of\nsubpopulations of plasmids derived from a single algal colony, which is consistent with\nwhat has been reported for conjugation-based delivery systems (see Diaz-Garza et al.,\n2024; doi: 10.1186/s12934-024-02559-y).\nEpigenetic silencing: We did not characterize the epigenetic fate of introduced episomes\nas it lies beyond the scope of our work. Furthermore, recent work from the Desgagné-\nPenix lab has examined the epigenetic fate of pCC1BAC-based episomes (same as in our\nstudies) introduced into P. tricornutum in detail (see Diamond et al, 2023; doi:\n10.1016/j.algal.2023.102998 and Diaz-Garza et al., 2024; doi: 10.1186/s12934-024-\n02559-y).\nNow, to the second comment:\n“Additionally, the novel and potentially impactful observation of in vivo DNA assembly in\nalgae is acknowledged but not explored in depth.”\nThis is again a very valid criticism, and we performed additional work investigating\npotential applications for DIVA as well as characterizing the parameters necessary for\nsuccessful assembly. Specifically, we have devised a system for reliably assembling any\ngene of interest into an episome directly in P. tricornutum and we also characterized that\noverlaps as small as 20 bp are sufficient for homology-directed assembly. We will delve\ninto further details regarding this below, but want to thank you for this feedback as it has\nstrengthened our work and opened endless avenues for exploration in the lab.\nBelow are specific comments and suggestions for revision:\nMajor comments:\nOverstatements\nThe authors claim that DNA delivery is the major challenge in diatom transformation. While\nthis is true for several diatom species, it does not hold for Phaeodactylum tricornutum,\nwhich can be efficiently transformed using multiple established methods. In contrast, a\nmore significant challenge in P. tricornutum is the frequent epigenetic silencing of\ntransgenes. Furthermore, P. tricornutum lacks a silica cell wall, making it particularly well-\nsuited for spheroplasting-based approaches.\nwithout proper context, and we have revised the text accordingly to better clarify the\nspecific challenges associated with DNA delivery in P. tricornutum.\nOur rationale for including this statement initially was that, prior to this work, the only well-\nestablished and widely used methods for transforming P. tricornutum were biolistics and\nconjugation. Although electroporation-based methods have been described since the early\n2010s, they were not widely adopted due to limited efficiency. Biolistics, while functional,\ntypically yields relatively few transformants and is not well suited for delivering large\nconstructs. Conjugation is more efficient but can have other problems. In our own\nexperience, some transmissible plasmids have proven incompatible with the canonical\nMob-based helper plasmids, necessitating extensive troubleshooting to achieve successful\nconjugation (this is the focus of our recent preprint – Walker et al., doi:\n10.1101/2025.10.16.682956). Moreover, some transgenes such as endonucleases (e.g.,\nCRISPR-Cas9, Cys4) can be toxic in E. coli, causing issues with plasmid stability prior to\nperforming conjugation.\nBy optimizing the electroporation method ~200-fold and developing a PEG-based\ntransformation protocol, we substantially expand the molecular toolbox for P. tricornutum\nand make this organism more accessible for genetic engineering.\nWe touched on this in our response above, but we did not investigate the epigenetic fate of\nintroduced episomes as this went beyond the scope of the work, which was focussed on\nexploring the limits of electroporation and PEG transformation methods. Recent work from\nthe Desgagné-Penix lab has investigated this using pCC1BAC-based episomes\n(mentioned in comment above).\nBut we agree that with the reviewer and we adjusted the text both in abstract and\nintroduction where were made such statements in our initial submission.\nThe inclusion of Thalassiosira pseudonana is also unconvincing. As the other main model\ndiatom species, T. pseudonana has a well-documented history of successful\ntransformations, and it is unclear how the current protocol represents a major\nadvancement for other species.\nWe appreciate the reviewer’s comment and the opportunity to clarify our rationale for\nincluding Thalassiosira pseudonana in this study. As noted in your comment above, P.\ntricornutum has a relatively poorly silicified cell wall, which may make it more amenable to\nspheroplasting/protoplasting transformation methods. We therefore sought to test whether\nour approach could also be applied to a more conventional diatom species with a siliceous\nfrustule.\nWe selected T. pseudonana because it already has well-characterized episomal systems\nand selectable markers, making it an ideal candidate for a proof-of-concept demonstration.\nWhile T. pseudonana can indeed be transformed using established biolistic and\nconjugation methods, the PEG-based approach we developed is considerably simpler and\nfaster to perform. As with P. tricornutum, this method expands the available molecular\ntoolbox and demonstrates that the protoplasting-based strategy can be extended to\ndiatoms with more heavily silicified cell walls.\nThe authors further propose that de novo diatom in vivo assembly (DIVA) could replace\n7\ntraditional cloning approaches in E. coli and S. cerevisiae, which they describe as time-\nconsuming. However, this claim is not well substantiated. The data presented show that in\nvivo assembly in diatoms is prone to errors, in contrast to the reliable transformation with\npre-assembled plasmids. Moreover, assembled episomes still require purification and\ntransformation into E. coli for verification. This statement also does not account for the\nlengthy selection process in diatoms, which typically takes around three weeks, compared\nto the single-day turnaround in E. coli. Instead of positioning DIVA as a general\nreplacement for bacterial or yeast cloning, we suggest that the authors propose a more\npractical and specific application where DIVA could offer a clear advantage.\nWe thank the Reviewers for this thoughtful comment. We agree that our original statement\n– that DIVA could broadly replace traditional cloning in E. coli or S. cerevisiae – was overly\ngeneral and not sufficiently supported by the data presented. As the Reviewers noted, in\nvivo assembly in P. tricornutum is more error-prone than cloning in bacteria or yeast, and\nassembled episomes must still be recovered and validated in E. coli. The longer selection\ntime in diatoms further limits the feasibility of positioning DIVA as an alternative for the\nstandard molecular cloning workflows.\nIn accordance with this comment, we performed additional experiments to better\ncharacterize both the limitations and potential applications of DIVA. To assess the minimal\nhomology requirements, we performed assembly using a NAT ORF amplified with 0, 20, or\n40 bp of overlap to the target episome; these experiments showed that 20 bp is sufficient\nto direct HDR, consistent with observations in S. cerevisiae. This result is practically useful\nas it means any gene or fragment of interest can be PCR-amplified with primers that add\n≥20 bp of overlaps and then directly used for DIVA. Previously, we relied on ~160 bp\noverlaps, which cannot be readily generated by PCR.\nWe next evaluated whether DIVA could be used to test putative localization signals. We\nfused a predicted mitochondrial signal peptide from the nuclear-encoded, mitochondrially\nlocalized protein CytB to eGFP and cotransformed this construct with an episome\ndesigned for insertion. Mitochondrial fluorescence was observed in 4 of 10 screened\ntransformants, supporting the functionality of the predicted CytB transit peptide.\nThe CytB-eGFP assembly was somewhat inefficient in that the episome could circularize\nthrough NHR, leading to nourseothricin resistance without the integration of the construct.\nTo address this, we synthesized another gBlock comprising eGFP linked to an incomplete\nNAT ORF via the previously characterized T2A linker. This design yielded 10/10\ntransformants with cytosolic eGFP expression upon screening, demonstrating that this\nstrategy provides a reliable approach for direct insertion and expression of a gene of\ninterest via DIVA.\nAt present, DIVA is viewed as a promising assembly capability rather than a fully optimized\nplatform. We established the basic parameters (e.g., overlap length, number of fragments,\nrelative efficiencies) and demonstrate two proof-of-concept applications. We anticipate that\nthe clearest advantages may emerge in contexts where rapid or pooled assembly directly\nin P. tricornutum is useful – such as screening promoter libraries or constructing variant\npools for phenotype-based enrichment – but exploring these applications lies beyond the\nscope of the present work. We have updated the Results section (Exploring the limits and\napplications of DIVA), added additional data (Supplementary Figs. 11 and 12), and have\nrevised the Abstract/Introduction/Discussion sections to present a more modest and\nbalanced interpretation of DIVA.\nplatform. We established the basic parameters (e.g., overlap length, number of fragments,\nrelative efficiencies) and demonstrate two proof-of-concept applications. We anticipate that\nthe clearest advantages may emerge in contexts where rapid or pooled assembly directly\nin P. tricornutum is useful – such as screening promoter libraries or constructing variant\npools for phenotype-based enrichment – but exploring these applications lies beyond the\nscope of the present work. We have updated the Results section (Exploring the limits and\napplications of DIVA), added additional data (Supplementary Figs. 11 and 12), and have\nrevised the Abstract/Introduction/Discussion sections to present a more modest and\nbalanced interpretation of DIVA.\nCo-transformation with RNP Cas9-gRNA complexes and episomal resistance\nmarker: The manuscript presents this strategy as a solution for targeting genomic loci that\nlack a selectable phenotype. However, with a reported co-transformation efficiency of only\n10%, the majority of resulting colonies (~90%) are false positives. The authors might\nconsider exploring an alternative strategy—such as co-delivery of multiple RNP complexes\ntargeting different loci—similar to the approach described in Serif et al., 2018.\nWe thank the Reviewers for this thoughtful comment and have added further clarification in\nthe revised manuscript to make the purpose of this experiment clearer (also in accordance\nwith comments made by Reviewer 1). Our aim was to quantify the probability of co-\ndelivering RNPs and episomal DNA into the same cell, facilitating future screens where\nCRISPR-induced genomic changes do not show an obvious phenotype/cannot be selected\nfor directly. Below we have an illustration to try to visualize what we were trying to quantify\n(Rudimentary Fig. 1); in short, a cell that takes up episomal DNA has some probability of\nalso taking up an RNP when transformed simultaneously.\nDetermining the co-delivery rate provides a useful benchmark for experiments with RNPs\ntargeting non-selectable loci as it allows us to estimate the number of colonies that should\nbe screened to identify edits. In this context, the ~10% co-transformation rate tells us that\nif we co-transform a selectable episome with an RNP targeting a non-selectable locus, we\nshould aim to screen approximately 10 colonies to identify 1 putative RNP-transformed\ncolony. Without this, we have no capacity to select for RNP-transformed cells when the\ngene being targeted doesn’t produce a selectable phenotype – a single transformation\nuses ~ 1 x 108 cells, after all. The 90% of transformants that didn’t demonstrate co-delivery\nof both the RNP/episome are not false positives in this case either; they are simply\ncolonies that were only transformed by one complex. We believe that our demonstrated\napproach will be highly useful for generating targeted deletions, including the potential\ncreation of a comprehensive deletion library of all non-essential genes.\nRudimentary Figure 1. When simultaneously electroporating an episome (i.e., DNA) and\nRNP into P. tricornutum, the vast majority of cells (i.e., >99.9%) will remain untransformed.\nOf the transformed cells, some will have taken up only the episome (red hemisphere) or\nthe RNP (blue hemisphere), with some proportion having taken up both constructs (purple\nintersection). We sought to characterize the percentage of transformed cells that took up\nboth the episome and RNP simultaneously to quantify the probability of co-transformation.\nExamples of overstatements:\npseudonana is also a model diatom species with a small genome, rapid growth, and\ngenerally accesible for genetic transformation. Given the differences between species and\nthat electroporation was only tested in P. tricornutum (and most effective on plates),\ncauction should be used when implying broad adaptability.\nWe thank the Reviewers for this helpful comment and have revised the text to use more\ncautious language regarding the adaptability of these methods. We had previously\nmentioned in the text that P. tricornutum is an atypical diatom with a relatively poorly\nsilicified cell wall, which may make these transformation techniques more amenable\ncompared to other species. We want to note that similar concerns were raised when the\nbacterial conjugation method was first established for P. tricornutum in 2015. Since then,\nconjugation – which also relies on direct DNA transfer – has been successfully adapted for\na range of diatom species, including T. pseudonana (as reported in the original 2015\nstudy), Fistulifera solaris, Nitzschia captiva, Cyclotella meneghiniana, and other species\nthat have yet to be published (anecdotal; based upon discussions during the MLD 8\nmeeting). We have added this context to the Discussion to support the rationale for\npotential adaptability of these methods. The text we added to the Discussion is as follows:\n“These results suggest that the spheroplasting and protoplasting techniques can be\napplied to other diatoms, though it will likely be necessary to optimize the intensity and\nduration of the enzyme treatment on a species-by-species basis along with other\nparameters in the respective protocols. This expectation aligns with the trajectory of\nbacterial conjugation in diatoms, which was originally established using plated P.\ntricornutum cells: although initially assumed to have limited applicability, conjugation has\nsince been successfully adapted to multiple species, including Cyclotella meneghiniana,\nNitzschia captiva, and Fistulifera solaris. This precedent underscores the realistic potential\nfor broader adaptability of the methods presented here.”\nLine 64: “Conjugation requires the creation and maintenance of a donor strain of\nEscherichia coli, which must contain a suitable broad-host-range conjugative plasmid in\naddition to the episome of interest.” While this statement is technically accurate, it is\nunclear why the authors present it as a drawback. The creation and maintenance of donor\nE. coli strains is a standard molecular biology procedure and typically not considered a\nmajor limitation.\nWe thank the Reviewers for this observation and have changed the text provide greater\ncontext for this statement. While creating donor E. coli strains is indeed a standard\nmolecular biology procedure, in practice it can present several unexpected challenges. As\nnoted above, we and others have encountered issues with incompatibility between certain\ntransmissible episomes and the MOB-based conjugative plasmids used for P. tricornutum\nand T. pseudonana (see our preprint – Walker et al., doi: 10.1101/2025.10.16.682956).\nFurthermore, co-maintaining MOB plasmids with medium- or high-copy transmissible\nplasmids can lead to rearrangements and plasmid instability, particularly when the\ntransmissible construct expresses genes that are considered toxic to E. coli (e.g.,\nendonucleases). Thus, when creating a conjugative E. coli strain, it’s best practice to\nreisolate and sequence the plasmid of interest after several passages to validate its\nstability and integrity over time. In contrast, direct transformation through electroporation or\nPEG transformation circumvents this step and can therefore shorten experimental\ntimelines by several days to weeks (and potentially even months, as was the case for our\n10\nplasmid incompatibility issue). Our intent here is to provide efficient alternative for DNA\ndelivery.\nWe have expanded the text in the Introduction section to elaborate on these practical\ndrawbacks. The improved text (in the Introduction section) now reads:\n“Conjugation requires the creation and maintenance of a donor strain of Escherichia coli,\nwhich must contain a suitable broad-host-range conjugative plasmid in addition to the\nepisome of interest; complications can arise if these two constructs have incompatible\nreplication machinery, if the episome is maintained at medium to high copy number, or if it\ncontains elements that are toxic when expressed in E. coli (e.g., endonucleases). These\nissues can lead to episomal rearrangements or loss over time. As a result, donor strains\noften require repeated reisolation and plasmid sequencing to confirm construct stability –\nsteps that can add days to weeks, and in some cases months, to a workflow.\nLine 89: Reiteration of line 28 comment: the claim that these methods can be broadly\napplied to other diatoms should be more cautious and supported with data or omitted.\nWe agree with this statement and have revised the text accordingly, as stated under the\nearlier comment regarding Line 28.\nGeneral note: The data suggest that spheroplasting improves transformation mainly on\nplates, with limited benefit in liquid culture. This is important because many diatom species\ncannot grow on solid media.\nWe thank the Reviewers for this important point. In our Discussion, we had previously\nstated that “Though P. tricornutum is unique in both its poorly silicified cell wall and\nwillingness to grow on solid media, we were able to adapt the alcalase-based\nprotoplasting technique for T. pseudonana…” to address this. The conjugation-based\nmethod, which also relies on plated cells for efficient P. tricornutum transformation, has\nbeen successfully adopted in several other diatom species, suggesting that plating\nrequirements are unlikely to represent a major barrier to broader adoption of\nelectroporation or PEG-based approaches. This ties into our response for the Line 28\ncomment above. We added the following text to better highlight our earlier statement:\n“Though P. tricornutum is unique in both its poorly silicified cell wall and willingness to\ngrow on solid media, we were able to adapt the alcalase-based protoplasting technique for\nT. pseudonana, a diatom species with a silicified cell wall that is typically cultured in\nliquid media.”\nUneven quality across the manuscript:\nThe final two sections—PEG transformation and Thalassiosira pseudonana\ntransformation—are notably underdeveloped compared to the rest of the experimental\nwork. These sections appear rushed, lacking sufficient detail, proper quantification, and\nclear presentation of results. To maintain consistency and clarity, the authors should either\nexpand and revise these sections to match the depth and quality of the earlier parts of the\nmanuscript or consider omitting them altogether.\nWe thank the reviewers for this valuable suggestion and have expanded both sections\naccordingly. The fluorescence microscopy of T. pseudonana PEG transformants was\nrepeated using proper controls and multiple channels to provide higher-quality and more\nconclusive data (Supplementary Fig. S17), and the P. tricornutum PEG section has been\nrevised to include additional methodological detail in the methods section. While the PEG\ntransformation was intended as a complementary proof-of-concept rather than a major\nfocus of the study, we agree that the additional data and clarification improve the overall\nconsistency and completeness of the manuscript.\nOpen questions:\nWould the addition of telomeric sequences help stabilize the linearized plasmids and\nprevent their circularization in diatom cells?\nThis is an excellent question, and coincidentally, this is (at least in part) where our\nexplorations began. We want to one day install a synthetic linear chromosome in the P.\ntricornutum nucleus as a part of the Pt syn 1.0 project (see Pampuch/Walker et al.; doi:\n10.1016/j.cogsc.2022.100611). As a first attempt at this, we PCR amplified the episome\npPtGE31_ΔPtR using primers that would add 0, 1, 2, or 5 telomere repeats (AACCCT) to\nthe termini. This was prior to the optimization of the electroporation method, so for these\ntransformations, we generated hundreds of transformants for each of the episomes tested\n(Rudimentary Fig. 2). Colonies from the 1T, 2T, and 5T transformation plates were picked\nand screened using a PCR primer set that would only generate an amplicon if the episome\nhad circularized (sparing any major deletions around the episomal termini). For 14 out of\nthe 15 colonies screened, an amplicon was generated – though the size of the amplicons\nvaried, which suggested to us that circularization was occurring, and indels were accruing\nduring circularization (Rudimentary Fig. 2). A few of the episomes were then recovered in\nE. coli, validating that circularization had indeed occurred (discussed further in response to\nReviewer #1, see Supplementary Fig. S2). We also sequenced four episomes, confirming\nthat the construct’s termini had been ligated through a non-homologous repair process.\nRudimentary Figure 2. Transformation of linear pPtGE31_ΔPtR with 0, 1, 2, or 5\ntelomere repeats at its termini. (a) pPtGE31_ΔPtR (~11 kb) was amplified using primers\nthat would add up to 5 telomere repeats at its termini. These constructs were transformed\nalongside the nourseothricin resistance marker (NAT) as a positive control, as up until this\npoint, we had not yet explored the delivery of episomes through electroporation (i.e., we\nhad only explored genomic integration of markers). (b) An agarose gel depicting the\namplicons generated from the constructs of panel a. An off-target band appears for the 5-\n12\ntelomere (5T) construct around the 2 kb mark. (c) Colonies post electroporation of the NAT\nmarker and various pPtGE31_ΔPtR telomere constructs. The negative control consisted of\ncells electroporated without episomal or integrative DNA.\nRudimentary Figure 3. Screening five transformants from the 1T, 2T, and 5T colonies\nshown in Rudimentary Fig. 2. DNA was isolated from the colonies and then subjected to\nPCR screening using a primer set spanning the junction termini. If the episomal ends were\nreattached seamlessly, we would expect a band of 267 bp (as demonstrated by the\npositive control, which was circular pPtGE31_ΔPtR). Any insertions or deletions could\nimpact the size or appearance of this amplicon. This screen was conducted using the\nSuperPlex kit (Takara; 35 cycles), with the negative control consisting of ddH O.\nknockout? What is the underlying metabolic or genetic rationale?\nWe appreciate the Reviewers’ question and have expanded the relevant section\naccordingly. As noted, this experiment was intended as a proof-of-concept based on the\nwork of Serif et al. (doi: s41467-018-06378-9), who characterized the APT/2-FA\ncounterselection system in P. tricornutum in detail. To our knowledge, PtAPT functions in\nadenine salvage rather than as an auxotrophic gene, and therefore adenine\nsupplementation is not strictly required for growth of PtAPT-knockout strains under normal\nconditions. However, we observed that adenine supplementation substantially improves\nrecovery of transformants following electroporation of the Cas9:PtAPT complex. When\nplated on 2-FA selection medium, plates lacking supplemental adenine produced only a\nhandful colonies, whereas plates with supplemental adenine yielded thousands\n(Supplementary Fig. S14).\nThis finding arose somewhat serendipitously. While optimizing 2-FA selection parameters\nin our hands (simply because different batches/manufacturers of 2-FA can vary in\npotency), wild-type P. tricornutum cells plated on media containing 5 mg/L adenine but no\n2-FA showed unexpectedly poor growth. Subsequent tests confirmed that adenine\nsupplementation of 1 to 5 mg/L was surprisingly toxic to wild-type cells (Supplementary\nFig. S13). Given this, we compared plating of Cas9:PtAPT transformants on 2-FA medium\nwith and without adenine to assess whether supplementation would affect recovery,\nhypothesizing that adenine might also stress PtAPT-knockout strains. Notably, Serif et al\nreported plating Cas9:PtAPT transformants on 2-FA plates supplemented with 5 mg/L\nadenine; we are uncertain why adenine was added in their experiments, it was not fully\n13\nclarified there either. In our experiments, as mentioned above, adenine supplementation\nproved to make a significant difference in the recovery of transformants (though our initial\nhypothesis was incorrect).\nMechanistically, APT catalyzes the transfer of phosphoribosyl pyrophosphate (PRPP) to\nadenine to form AMP. In the presence of 2-fluoroadenine (2-FA), APT instead converts 2-\nFA into 2-fluoro-AMP, which is subsequently phosphorylated to form toxic fluorinated\nnucleotides (i.e., 2-fluoro-ADP/ATP). When APT is knocked out, this conversion no longer\noccurs, preventing toxicity. Based on this, we are uncertain why the addition of adenine\nimproves the recovery of PtAPT transformants so drastically; perhaps there are other\nadenine salvage/recycling/metabolic pathways, and the presence of supplemental adenine\nreduces the toxicity of 2-FA in these alternative pathways. This would be our best\nhypothesis, especially since adenine supplementation is not necessary when passaging\nthe cultures in the absence of 2-FA.\nTogether, these results suggest that while adenine supplementation is not strictly required\nfor PtAPT-knockout strains, it enhances transformant recovery and acts as a metabolic\nbuffer under 2-FA selection. The adenine toxicity observed in wild-type cultures further\nimplies that purine salvage and PRPP homeostasis are tightly regulated in P. tricornutum.\nFurther investigation into this regulation would be valuable but lies beyond the scope of\nour work; we have added some text to comment on this phenomenon in the Results\nsection and invite further inquiry in the Discussion section. The text reads as follows:\nRESULTS\n“Serif et al. had previously shown that knock-out of the P. tricornutum Adenine\nPhosphoribosyl Transferase (PtAPT) gene confers resistance to the cytotoxin 2-\nfluoroadenine (2-FA) at a dose of 10 µM. PtAPT functions in the adenine salvage pathway,\ncatalysing the transfer of phosphoribosyl pyrophosphate (PRPP) to adenine to form\nadenosine monophosphate (AMP); in the presence of 2-FA, PtAPT instead converts 2-FA\ninto 2-fluoro-AMP, which is subsequently phosphorylated to form toxic fluorinated\nnucleotides.\nChemicals can vary based on the manufacturer and/or batch; thus, when setting forth on\nthis series of experiments, we first tested the resistance of wild-type P. tricornutum (CCAP\n1055/1) cultured on ½-salt L1 plates supplemented with adenine (5 µg/ml) and our batch of\n2-FA (0, 2.5, 5, 10, 15, 20 µM). To our surprise, there was an absence of growth on the 0\nµM 2-FA plate, suggesting that adenine alone posed a toxicity to wild-type P. tricornutum\ncells. We tested this theory by plating wild-type cells onto ½-salt L1 plates supplemented\nwith varying levels of adenine (0, 1, 2.5, 5 µg/ml; Figure S13). The growth of wild-type\nalgae was negligible after one-week on the 5 µg/ml adenine plate, but relatively\nunimpacted on the 0.5 and 1 µg/ml plates. This interesting revelation led us to hypothesize\nthat adenine supplementation could negatively impact ΔPtAPT transformants as well, so\nwhen conducting our RNP electroporation experiments, we plated transformants on 2-FA\nplates with and without adenine supplementation.\n…\nTransformants were recovered for one day before plating one-fifth of the reaction across ½\nsalt L1 plates supplemented with 10 µM or 5 µM 2-FA, with or without 5 µg/ml adenine\n(Supplemental Fig. S14). This pilot experiment generated thousands of 2-FA resistant\ntransformants across the plates that had been supplemented with adenine, but only a\n14\nhandful across the plates without supplemental adenine. This was an unexpected\ndiscovery given our previous observations. Ten 2-FA resistant colonies were passaged for\nfurther screening at the PtAPT loci to determine if Cas9-mediated engineering had\noccurred.”\nDISCUSSION\n“Through this series of proof-of-concept experiments, we also uncovered an unexpected\nparadox: while adenine supplementation (5 µg/ml) is visibly toxic to wild-type P.\ntricornutum, it substantially improves the recovery of ΔPtAPT transformants under 2-FA\nselection. Because PtAPT functions in the adenine salvage pathway, it is not immediately\nclear why supplemental adenine would have any beneficial effect on ΔPtAPT strains; if\nanything, one might expect the opposite. This result suggests that additional adenine\nsalvage, recycling, or purine-metabolic pathways may be active in P. tricornutum, and that\nsupplemental adenine may mitigate 2-FA toxicity through these alternative routes.\nElucidating the underlying metabolic regulation will require further investigation, but is\nbeyond the scope of the present study.”\nAdditional line-by-line comments:\nLine 26: The term “PEG” is used in the Abstract without being defined. Please define\n“polyethylene glycol” at first mention.\nThank you for catching this, we have corrected the Abstract accordingly.\nLine 63: The transformation rate is incorrectly written as “4 transformants per 10⁻⁴ cells”. It\nshould read “4 transformants per 10⁴ cells” for consistency with line 58.\nThank you for catching this, we have corrected this error and all other related errors\naccordingly.\nLine 69: More background and context on PEG-mediated transformation in diatoms is\nneeded. Currently, the discussion is too brief for readers unfamiliar with the technique.\nWe appreciate this suggestion and have expanded the background on PEG-mediated\ntransformation accordingly. To our knowledge, PEG transformation has only been\ndescribed once in diatoms (Karas et al., doi: 10.1038/ncomms7925), where the method\nwas reported to be inefficient and inconsistent, yielding only 2–3 transformants across\ndozens of attempts. In revisiting this approach, we followed the overall workflow of Karas\net al. but introduced several key modifications: we used alcalase as the protoplasting\nenzyme instead of the previously reported zymolyase/hemicellulase/lysozyme cocktail, we\nincluded salmon sperm DNA during the transformation step, and we recovered the\nprotoplasted cells in liquid media and for a longer duration before plating. These\nadjustments substantially improved the robustness and reliability of the protocol, routinely\nproducing thousands of transformants per experiment when transforming pPtGE31_ΔPtR.\nWe have added the following text to the background section:\n“Although PEG-mediated transformation has not yet been demonstrated in other diatom\nspecies, it is routinely applied in the microalga Chlamydomonas reinhardtii, where cell-wall\nremoval followed by PEG-mediated agitation with DNA and glass beads enables efficient\nnuclear transformation.”\nThank you for catching this, we have corrected this error.\nLine 87: The manuscript inconsistently uses the terms spheroplasting and protoplasting.\nThese are sometimes used interchangeably and sometimes as distinct processes. Given\nthe title refers specifically to “spheroblasted cells,” a clear and consistent definition is\nessential.\nWe thank the Reviewers for noting this inconsistency. We have reviewed the manuscript to\nensure the terminology is used consistently and now clearly define our usage of\n“spheroplasting” and “protoplasting”. The following text was added to make this distinction\nclear:\n“Notably, a spheroplasted cell has only partial cell wall degradation and may not appear\nphenotypically different from a wild-type cell, whereas a fully protoplasted cell lacks a cell\nwall entirely and adopts a characteristically spherical morphology – provided it does not\nburst from osmotic or mechanical stressors.”\nWe also propose a change the title of the manuscript to read “Breaking the cell wall: a\ngamechanger for DNA delivery to diatoms” to hopefully resolve this inconsistency.\nLine 97: The authors note that a neighbouring lab used older electroporation parameters\nand that the switch from 25 to 50 µF may relate to the equipment used. This raises the\nquestion of whether the improved transformation efficiency is protocol-specific or simply\nequipment-dependent. Please clarify.\nWe appreciate the Reviewer’s point and want to clarify – the substantial increase in\ntransformation efficiency results from the spheroplasting step, not from changes to the\nelectroporation pulse parameters. Prior to optimizing the resistance settings for our\nspecific electroporator, we were unable to obtain transformants using the previously\nreported parameters (i.e., zero transformants across tens of experiments); after adjusting\nthe resistance to values compatible with our equipment, we achieved baseline\ntransformation efficiencies of ~10-250 colonies per experiment. It was the addition of the\nspheroplasting step that produced the large, order-of-magnitude improvement reported in\nthe manuscript.\nOur note about a neighbouring lab successfully using the original pulse settings was\nintended only to emphasize that electroporation parameters can be equipment-dependent,\nand users may need to empirically optimize these settings for their own systems. The\nunderlying boost in efficiency is protocol-specific and reflects the impact of spheroplasting,\nnot differences in electroporators.\nLine 101: Plasmid pPtGE31_ΔPtR with ID 236260 is not found on Addgene. Please upload\nthe plasmid map and sequence before any resubmission.\nAll of our plasmids had been submitted to Addgene at the time of initially submitting this\nmanuscript, they can only be found by typing the ID directly into the link since they are not\npublicly available yet. These plasmids will be released upon publication of the manuscript.\nthe last 6 numbers in the link with the corresponding ID: https://www.addgene.org/236260/.\nLine 176: The use of 500 ng of a PCR-amplified 11-kb construct should be mentioned\nearlier in the text, especially when referring to Figure 1.\nWe agree and have made the corresponding change.\nLine 256: The claim that linear pSC5 is three times more efficient than circular is not\nclearly supported by Figure 3B, where colony counts appear similar (5 vs. 6 CFUs). Please\nclarify.\nWe agree and have removed this statement, the colony counts are not substantially\ndifferent enough to support this claim.\nLine 302: Plasmid pPtGE31_ShBle with ID 236261 is not found on Addgene. Please\nupload the plasmid map and sequence before any resubmission.\nSee the response to the comment on line 101 – you can find the plasmid here:\nhttps://www.addgene.org/236261/. It will not be searchable or publicly available until the\nmanuscript is published. Everything has been sequence validated by Addgene and is\nready for distribution otherwise.\nLine 393: There is no Figure S6C. This likely refers to Figure S7C—please correct.\nThank you for catching this, we have corrected this in the text.\nwould strengthen the end of the manuscript.\nThis is an excellent suggestion. This was also asked by first reviewer and we added the\nfollowing text:\n\"The aim of this experiment was to develop protocol that could significantly improve\nscreening for gene deletions that do not produce an obvious phenotype.”\nLine 550: The difference in DNA quantities used (500 ng linear vs. 25 µg circular) is\nsubstantial. Please explain how these values were chosen.\nWhen this method was first explored by Karas et al in the 2010s, PEG transformation\nefficiency was very low (2-3 colonies across several experiments), and it was standard\npractice to add as much circular plasmid DNA as possible in an attempt to increase the\nnumber of transformants. At that time, it was not yet known that linearized episomal DNA\ntransforms more efficiently than circular DNA in diatoms.\nIn revisiting the protocol for the present study, we intentionally used 500 ng of linear\nepisomal DNA (pPtGE31_ΔPtR) so that the PEG-based transformations could be directly\ncompared to our electroporation experiments, all of which also used 500 ng of the same\nconstruct. The fact that we were able to obtain thousands of transformants with only 500\nng of DNA further highlights the substantial improvements in efficiency achieved with the\nupdated PEG protocol.\nLine 554: Early in the manuscript, alcalase dosage is reported in mAnson units per 10⁸\ncells, but here it is given in µl. Please standardize units throughout.\nWe thank the Reviewers for noting this inconsistency. We have now added the\ncorresponding mAnson unit values wherever alcalase dosage is reported.\nLine 558: The text states that 40 µg of carrier DNA improved electroporation efficiency\n~15-fold, but no results are shown for PEG transformation. Was this amount used in PEG\nprotocols? Was a similar improvement observed?\nWe also tested the addition of carrier DNA in the PEG-based transformations\n(Supplemental Fig. S15B, condition #1 vs #2) and reported ~10-fold difference, similar to\nwhat is seen for electroporation. For this step, the key factor is providing sufficient\nnonspecific DNA to protect the episomal construct from DNases secreted or released by\nP. tricornutum cells; in our experience, 40 µg is more than adequate (especially when\nusing 500 ng of an episome, ssssDNA is 80-fold more abundant).\nLine 565: The final optimized PEG protocol is not clearly presented. Please summarize the\nfinal conditions used and their effectiveness.\nThank you for this comment, we have revised the text to provide a more clear description\nof the PEG protocol.\nLine 566: There is no Figure S2D, despite the reference. Please correct or include the\nfigure.\nLines 565–571: This paragraph lacks quantitative data and clarity. It is difficult to assess\nthe advantage of PEG transformation without specific efficiency data.\nWe thank the Reviewers for this comment. We did not perform enough biological or\ntechnical replicates to generate quantitative data for the PEG approach (i.e., lacking data\nthat can be statistically analysed). The PEG protocol was included in this manuscript to\ndemonstrate a robust, equipment-independent alternative for DNA delivery – particularly\nfor species where electroporation may be challenging because the pulse parameters have\nyet to be quantified.\nAlthough we do not provide formal efficiency measurements, the PEG method consistently\nproduced readily recoverable numbers of transformants under the optimized conditions\ndescribed (using 500 ng linear pPtGE31_ΔPtR – 1000s of colonies generated), and thus\nrepresents a practical, reproducible alternative approach. The plates in Supplemental Fig.\n15C conditions #2 to 5 demonstrate the efficiency, keeping in mind that only ¼ of the\ntransformants are shown on these plates. We have revised the text to clarify the purpose\nof this section accordingly.\n“A polyethylene glycol (PEG) transformation method for P. tricornutum with an efficiency of\n≤ 1 transformant per 108 cells was previously described by Karas et al. For this method,\nalgal cells were protoplasted using a combination of zymolyase, lysozyme, and\nhemicellulase ahead of PEG treatment. We revisited this method using alcalase as the\nsole enzyme for cell wall removal, with the intention of providing a simple and equipment-\nindependent alternative to electroporation that is particularly useful for labs or species\nwhere this method remains a practical barrier.”\nLine 574: Reiterate the need to clearly outline the final optimized conditions.\nWe have resolved this in accordance with the comment on Line 565.\nLine 577: Please include the size of the episome used here, as done in the Materials and\nMethods section.\nThank you for catching this, we have included the size of pBIG1 (8224 bp, listed as ~8.2\nkb).\nLines 578–581: The results described are vague. Clarify what “some degree of growth”\nmeans and how harsher protoplasting conditions influenced colony formation.\nThis is a valid point. In the revised manuscript, we now refer specifically to the emergence\nof colonies across the different alcalase treatments. Because T. pseudonana does not\ngrow robustly on solid media, reliable colony counts were difficult to obtain for the various\nconditions. Although transformed T. pseudonana cells do form discrete colonies, we often\nobserved smeared or diffuse growth which made accurate counting challenging. We have\nrevised the text to make this more clear.\nLine 588: The phrase “a higher proportion of protoplasted cells are visible…” is vague.\nProvide more concrete observations or quantification, if available.\nfollowing treatment with 10 or 1000 ul of alcalase (a snapshot is presented in\nSupplemental Fig. S16; proper quantification was not performed). Based on these images,\nit appears as though ~ 1/28 visible cells are protoplasted in panel b (= 10 ul alcalase),\nand ~ 13/20 are protoplasted in panel c (=1000 ul alcalase). We have added this to the\ntext to provide a rough quantification of the extent of protoplasting with different alcalase\namounts. We have added the following text to address this:\n“Based on these microscopy images, cells from the 10 µl treatment showed a protoplasting\nrate of ~4% (1/28 cells), whereas cells from the 1000 µl treatment demonstrated a\nprotoplasting rate of ~50% (13/20 cells).”\nLine 593: There is no discussion on partial vs. complete cell wall removal. Is this\ndependent on enzyme concentration? Do cells survive full wall removal?\nFor the PEG method, we observed that higher amounts of alcalase and longer incubation\ntimes (i.e., more protoplasted cells) yielded more transformants for both diatom species\ntested. Fully protoplasted P. tricornutum and T. pseudonana cells survive complete cell\nwall removal, but care must be taken during handling, as the cells are more fragile and\nprone to mechanical rupture. It is also important to maintain cells in full-salt L1 (or F/2)\nmedia to prevent bursting due to osmotic pressure. In contrast, for electroporation,\npartially spheroplasted cells can be susceptible to bursting because they are washed in\n375 mM D-sorbitol, which has lower osmolarity than L1; this issue is avoided in the PEG\nmethod, as cells are maintained in L1 throughout the procedure. We have made changes to the PEG\nsection (in accordance with prior suggestions) to highlight this and to state that >80% of\nthe P. tricornutum cells should be protoplasted before proceeding to the next step.\nLine 611: It is true that plated cells yielded the highest transformation rates, but based on\nTable S2, efficiencies are mostly in the 10⁶–10⁷ range, with only treated plated cells\nshowing higher values (1.19 x 10⁴). The benefit of alcalase treatment seems to apply\nmainly to plated cells.\nThis interpretation is correct based on the experiments we conducted; however, we did not\npursue extensive optimization of liquid-grown cultures because plate-derived cells were\nalready sufficient for establishing a robust protocol in P. tricornutum. Numerous variables\nremain unexplored in liquid cultures, for example recovery in liquid rather than on plates,\nor selection conditions better suited to cells not adapted to solid media, time of\nprotoplasting and amount of alcalase. It will be interesting to determine how many\nadditional diatom species benefit from cell wall removal, but we do not believe P.\ntricornutum is unique in this regard.\nLine 612: The manuscript states that transformation is “efficient,” but no numerical\nthreshold is provided. Please define what constitutes efficiency in this context.\nThank you for pointing this out, we have added the previously stated efficiency (~6\ntransformants per 106 cells) to this sentence to make it more clear.\nLine 645: “Several-fold more efficient” should be quantified.\nresistant transformants across their two biolistic experiments with RNPs targeting the\nPtAPT locus. We generated >1000 transformants in a single experiment (it was difficult to\ncount given the size of the colonies, but given that we plated 1/5 of the reaction per plate,\nthis number is likely closer to ~5000 per transformation), so we will report this as a >200-\nfold increase to be conservative.\nLine 701: Only PEG transformation was adapted to T. pseudonana. The claim that these\nmethods hold great promise for other diatoms lacks sufficient supporting evidence.\nAs noted in earlier responses, while we only tested PEG transformation in T. pseudonana,\nthe successful adaptation of this method to a diatom with a canonical silicified cell wall\ndemonstrates its broader potential. Similar hesitancies were raised when bacterial\nconjugation was first developed for P. tricornutum; yet, this approach has been\nsuccessfully adopted across a diverse range of diatom species. We have revised the text\nin accordance with an earlier comment (Line 28) to clarify this context and temper claims\nregarding broad applicability while highlighting the proof-of-concept value of our results.\nLine 866: The note about keeping materials on ice is useful and should be moved earlier in\nthe protocol section for clarity.\nThis is a great suggestion, we have added this step earlier to the protocol in accordance.\nLine 880: The DNA input range (“1 to 1000 ng”) is very broad. Consider adding a note that\nfor ~11-kb episomes, 500 ng appears to saturate the reaction.\nWe agree with this and have added a note to the methods section in accordance.\nLine 904: This line again references equipment differences affecting electroporation\nsettings. Consider addressing this earlier in the Methods for better context.\nWe agree with this and have accordingly positioned this text at the start of this\nmethodology section.\nFigure Comments:\nFigure 1b: Why is the image shown with 100 mAnson if the best treatment was achieved\nwith 1 mAnson? This is confusing and needs explanation.\nWe wanted to depict what over spheroplasted/protoplasted cells look like; when the cells\nare spheroplasted with 1 mAnson, they do not look different from untreated cells. In\naccordance with suggestions from Reviewer 1, we have added high resolution microscopy\nimages for the 0.1 mAnson, 1 mAnson, and 10 mAnson treatments to further characterize\nthe visible changes that emerge during the different spheroplasting intensities\n(Supplemental Fig. S1).\nFigure 1e and f: Statistical analysis should be included. Please clarify what alcalase\ntreatment was used and provide full experimental conditions (e.g., DNA concentration,\ntreatment duration) so that the figure is self-contained.\nThank you for this comment, we have now performed statistical analyses to demonstrate\nthat alcalase-treated, plated cells show significantly higher electroporation efficiency, as\npresented in Fig. 1E. Statistical testing was not applied to Fig. 1F, as this panel was\nintended to illustrate qualitative trends rather than compare the different DNA\nconcentrations quantitatively. We have also updated the Fig. 1 legend to provide additional\ndetails on the experimental conditions for panels E and F\nFigure 1e: As noted above, experimental details (e.g., DNA concentration, alcalase\nprotocol) are missing. These are essential for interpretation.\nWe have addressed this in accordance with the comment above, thank you for catching\nthis.\nFigure 1f: Clarify whether this experiment was done in liquid culture or on plates, and\nspecify the alcalase treatment used.\nWe have addressed this in accordance with the comment above, thank you for catching\nthis.\nFigure 2c–e: Please clarify the annotations in panels d and e to clearly indicate which PCR\nprimer pairs were used, with explicit reference to panel c.\nThank you for pointing this out, we have made the changes to Fig. 2 accordingly.\nFigure 3e is not clearly explained. Presumably, it should refer to line 269: “conjugation was\nsuccessful for seven out of eight E. coli pools”. If it is the case, then labelling C1-C4 in both\ncases is confusing.\nWe have revised Fig. 3 and the corresponding in-text reference to make this clearer, thank\nyou for bringing this to our attention. To us, it makes the most sense to keep the labelling\nas 1 through 4 as we were comparing two conditions (linear vs circular pSC5); to make\nthis distinction clearer to the reader, however, we relabelled the linear-pSC5-derived\ncolonies with L1 to L4.\nFigure 5: Part a shows three schemes of possible assembly. Part b does not correspond to\npart a. Please adjust for clarity.\nThank you for this helpful comment, we agree that the distinction between parts A and B/C\ncould be made clearer. Our intention in the current version of Fig. 5 was to present the\nepisomal assembly workflow in a way that captures the full process: an overview\nschematic of the possible assembly configurations, followed by one representative plate\nseries, and then one representative set of screening gels. This mixed presentation allows\nthe reader to see both the experimental outcome (colonies) and the downstream\nverification (screening PCRs in P. tricornutum and E. coli), rather than limiting the figure to\nonly plates or only gels.\nWith that said, we recognize that this layout may not appear directly matched to the\nschematics in part A. To address this, we have prepared two alternative versions of Fig. 5\n– one showing all plate series, and one showing all screening gels – to more closely mirror\nthe structure of panel (a). If the Reviewers feel that either alternative improves the clarity of\nthe Figure/manuscript, we are willing to change it to what they see as best fit.\nall of the screening gels are presented below the assembly schematic.\nHere, all of the experimental plates are presented below the assembly schematic.\nReviewer #3 (Remarks to the Author):\nI co-reviewed this manuscript with one of the reviewers who provided the listed reports.\nThis is part of the Nature Communications initiative to facilitate training in peer review and\nto provide appropriate recognition for Early Career Researchers who co-review\nmanuscripts.\n– your comments have greatly strengthened our work.\n25"}, {"round": 2, "reviewer_comments": {"reviewer_1": "The authors have addressed my peer review comments appropriately. I have no further points to raise.", "reviewer_2": "We would like to thank the authors for carefully addressing our previous comments. The manuscript is now substantially improved, and we particularly appreciate the more in-depth analysis of in vivo assembly in diatoms, which opens exciting new avenues for future research. While we have one major comment and a few minor remarks, we consider this work a valuable resource for the diatom research community and believe it deserves publication in Nature Communications.\nMain comment\nWhile the work on Phaeodactylum tricornutum is very convincing, the transformation of Thalassiosira pseudonana still raises some questions.\nThe circular episome pBIG1 (8.2 kb) conferring nourseothricin resistance and carrying eGFP was used in these experiments. \n– Is the sequence of this plasmid available or will it be made available?\n – Which promoter drives the expression?\n– Does it express only eGFP, or are there other expression elements present?\nAlthough in most cases the alcase units were standardized to mAnson, in this part off the manuscript, alcalase is still in μl alcalase\nIn Supplementary Fig. S17, the statement “Both autofluorescence and eGFP signals were visible in the pooled transformed cells, demonstrating that episome transfer had occurred” is difficult to evaluate. The figure is challenging to interpret:\nalthough there is no GFP signal in the untransformed control, the green signal observed in the transformed sample appears in structures lacking chloroplast autofluorescence and not corresponding to T. pseudonana cells. This raises concerns about what is being visualized and whether the image may be overexposed.\nMinor remarks\n- Although the authors have corrected most instances of “μl” to proper enzyme activity units for alcalase, a few remain, mainly in the T. pseudonana section.\n- Line 80: “Electroporation has a higher transformation efficiency (2.6 to 4.5 transformants per 10⁵ cells⁵,⁷)” → Please standardize the citation format with line 62, either inside or outside the parentheses.\n- Line 354: The correct reference is Supplementary Table S6, not S5.\n- Line 635: “develop a protocol” → missing the article “a”.\n- Keep the rudimentary Figure 5 with the plates in the main paper, and move the figure with the gels to the Supplementary Information.\nSide remark (for future consideration):\nIt is possible that the 5× telomeric repeat used is not sufficient to establish a fully protective protein complex equivalent to the human shelterin complex. For future experiments, the authors might consider testing longer telomeric repeats, although these can be difficult to clone and maintain in bacteria.", "reviewer_3": "I co-reviewed this manuscript with one of the reviewers who provided the listed reports. This is part of the Nature Communications initiative to facilitate training in peer review and to provide appropriate recognition for Early Career Researchers who co-review manuscripts."}, "author_response": "RESPONSE TO REVIEWERS\nOur responses are in purple.\nIn addition to the comments left by the reviewers, we identified a mistake in our last submission – we had cited ASAFind as the resource we used to identify the CytB signal peptide, when in fact, we had used TargetP (version 2.0). ASAFind was used in a separate project to identify bipartite chloroplast targeting signals of putative nuclear-encoded chloroplast-localized proteins of P. tricornutum. The canonical mitochondrial localization signal in diatoms is unipartite and shares conserved features across eukaryotes; thus, it can be identified using TargetP or SignalP. This has now been corrected in the manuscript.\nWe have also made changes in accordance with the NCOMMS Author Checklist (e.g., refining the abstract, reordering of manuscript sections, changes to word choices).\nReviewer #1 (Remarks to the Author):\nThe authors have addressed my peer review comments appropriately. I have no further points to raise.\nWe thank Reviewer 1 for their constructive comments during the initial review and are pleased that the revised manuscript has addressed their concerns.\nReviewer #2 (Remarks to the Author):\nWe would like to thank the authors for carefully addressing our previous comments. The manuscript is now substantially improved, and we particularly appreciate the more in-depth analysis of in vivo assembly in diatoms, which opens exciting new avenues for future research. While we have one major comment and a few minor remarks, we consider this work a valuable resource for the diatom research community and believe it deserves publication in Nature Communications.\nWe thank Reviewers 2 and 3 for their careful re-evaluation of the revised manuscript and for their encouraging assessment. We will address the remaining major and minor comments below.\nMain comment\nWhile the work on Phaeodactylum tricornutum is very convincing, the transformation of Thalassiosira pseudonana still raises some questions.\nThe circular episome pBIG1 (8.2 kb) conferring nourseothricin resistance and carrying eGFP was used in these experiments.\n– Is the sequence of this plasmid available or will it be made available?\nYes, the plasmid sequence will be made publicly available. This plasmid is also used in a separate manuscript that is currently under review (Mock lab), and we plan to release the full sequence upon finalization of that work. It will be deposited to Addgene as we have done in the past. Please see link for our lab materials deposited at Addgene: https://www.addgene.org/Thomas_Mock/.\n– Which promoter drives the expression?\nThe promoter is FCP (fucoxanthin-chlorophyll a/c binding protein), which is a common promoter used in diatom Thalassiosira pseudonana.\n– Does it express only eGFP, or are there other expression elements present?\nThe plasmid has also a T. pseudonana gene of unknown function (NCBI ID: XP_002293646) as this plasmid is being used for more than one project. However, this does not impact the expression of eGFP, which is the purpose of this study.\nAlthough in most cases the alcase units were standardized to mAnson, in this part off the manuscript, alcalase is still in μl alcalase\nThank you for catching this, we have standardized the alcalase units in this section and in Supplementary Fig. S16 to be consistent with the rest of the manuscript.\nIn Supplementary Fig. S17, the statement “Both autofluorescence and eGFP signals were visible in the pooled transformed cells, demonstrating that episome transfer had occurred” is difficult to evaluate. The figure is challenging to interpret: although there is no GFP signal in the untransformed control, the green signal observed in the transformed sample appears in structures lacking chloroplast autofluorescence and not corresponding to T. pseudonana cells. This raises concerns about what is being visualized and whether the image may be overexposed.\nHope we can address these concerns by explaining how microscope images of T. pseudonana are being influenced by the morphology of the cells. Their morphology is very different to P. tricornutum and if the reviewer is not so much familiar with T. pseudonana, it is likely that images are being misinterpreted.\nMorphology: Generally, T, pseudonana cells under the microscope have two different optical axes according to their orientation (Rudimentary Fig. 1). One is called valve view, and the other one is called girdle-band view. If the cells are imaged in valve view, they appear as round discs as shown in our paper. Those round discs might be misinterpreted as not being cells of T. pseudonana. However, they are the same cells but just in valve view.\nRudimentary Figure 1. Thalassiosira pseudonana under transmission electron microscope. (https://asknature.org/strategy/protein-directs-silica-growth/attachment/thalassiosira-pseudonana/)\nChlorophyll autofluorescence: If cells are imaged in valve view, it is much more challenging to obtain a strong chlorophyll signal because of the cellular content pilling up vertically and if chloroplasts are at the bottom of the pile, the signal of chlorophyll autofluorescence will be very weak (depending on orientation and focal depth), which is what we see sometimes in the image we have added to the paper. Thus, weak autofluorescence does not indicate absence of the cell especially because the cell outline was confirmed by the brightfield channel of the microscope.\nExposure / saturation control: The fluorescence images were not overexposed during acquisition. We ensured that camera settings avoided signal clipping (no saturated pixels) and that the transformed and untransformed samples were imaged using the same acquisition settings. To further alleviate concern, we provide an additional representative field of view (same strain, same imaging session) in which the cell morphology and the two fluorescence channels are clearer (Rudimentary Fig. 2).\nA) B)\nC) D)\nRudimentary Figure 2 T. pseudonana transformed cells under fluorescence microscope. A) for bright field, B) for merge, C) for chloroplast autofluorescence and D) for eGFP signal.\nMinor remarks\n- Although the authors have corrected most instances of “μl” to proper enzyme activity units for alcalase, a few remain, mainly in the T. pseudonana section.\nThank you for catching this, we have gone through the manuscript again and have hopefully caught all remaining instances of this. In many sections, we report both the enzyme activity (Anson units) and the corresponding volume (µl). Our reasoning for doing so is to illustrate how ‘potent’ the enzyme is/how little was needed for spheroplasting in the electroporation section (1 µl of enzyme per ~ 3 x 108 cells!). If preferred, we are happy to standardize all reporting to Anson units alone.\n- Line 80: “Electroporation has a higher transformation efficiency (2.6 to 4.5 transformants per 10⁵ cells⁵,⁷)” → Please standardize the citation format with line 62, either inside or outside the parentheses.\nThank you for catching this, we have placed the citations outside of the bracket and have scanned the manuscript for any other instances where we inconsistently formatted the citations.\n- Line 354: The correct reference is Supplementary Table S6, not S5.\nThank you for catching this, we have corrected the Table reference accordingly.\n- Line 635: “develop a protocol” → missing the article “a”.\nThank you for catching this, we have corrected the typo in the manuscript.\n- Keep the rudimentary Figure 5 with the plates in the main paper, and move the figure with the gels to the Supplementary Information.\nWe agree with this suggestion and have made the according changes to the manuscript and supplementary file. We have also updated the in-text references to correspond with these changes accordingly.\nSide remark (for future consideration):\nIt is possible that the 5× telomeric repeat used is not sufficient to establish a fully protective protein complex equivalent to the human shelterin complex. For future experiments, the authors might consider testing longer telomeric repeats, although these can be difficult to clone and maintain in bacteria.\nYes, this is a great point. We previously did a bioinformatic analysis of the characterized telomeres in P. tricornutum and saw that there are as few as 11 repeats naturally, which is remarkably ‘short’ compared to human telomeres. Still, it will likely be necessary to increase telomere/subtelomere length beyond a mere 5 repeats for stable long-term maintenance. Another hypothesis that we are contending with is that our initial construct may have simply been too small (~12 kb). Previous work done in S. cerevisiae demonstrated that linear chromosomes smaller than 100 kb cannot be stably replicated unless circularized – presumably due to issues with replication and/or segregation. We have work underway to try and characterize these parameters as our ultimate goal is to establish synthetic chromosomes in P. tricornutum (as a part of the Pt syn 1.0 project).\nReviewer #3 (Remarks to the Author):\nI co-reviewed this manuscript with one of the reviewers who provided the listed reports. This is part of the Nature Communications initiative to facilitate training in peer review and to provide appropriate recognition for Early Career Researchers who co-review manuscripts."}]

Decision letter