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Diversity and inclusivity
From previous research, we were inspired that bacterial communities can affect the formation and outbreak of green tide.
Among them, six bacterial functional groups (BFGs)[1] are defined. In the early stage of green tide migration, growth regulating BFG I and antibacterial stress resistant BFG II dominate, which can regulate algal growth and synergistically protect it; In the later stage, heterotrophic BFG III and algicidal BFG IV dominate, competing with algae for nutrients and inhibiting their growth; Nutritious BFG V and algal derived trophic BFG VI coexist with algal hosts. By using high-throughput sequencing technology of 16S rRNA and clustering analysis of operational taxonomic units (OTUs)[2], the bacterial community structure was determined, and the roles of different bacterial functional groups in green tide migration were analyzed, revealing the spatiotemporal complexity of the interaction between bacteria and Ulva prolifera.
Fig1. Potential algal-bacteria interactions during the migration stages of the U. prolifera green tide.
Inspired by that, we chose bacterial biofilm (a thin, sticky layer formed by groups of bacteria that cling to surfaces and are held together by a slimy substance they secrete) to capture Ulva spores, partly because biofilm has a large surface area and special microstructure, which can provide more attachment sites for Ulva spores and increase the probability of capturing spores; On the other hand, microorganisms and their secreted substances in the biofilm may have an attractive effect on the spores of Ulva prolifera, and can create a relatively stable microenvironment that is conducive to the attachment and survival of spores, which is helpful for further research on the reproduction of Ulva prolifera and the formation mechanism of green tide.
Since Ulva prolifera and its related research belong to a niche field, all the literature we collected focuses on the genus Ulva rather than the specific species Ulva prolifera—primarily due to the scarcity of species-specific studies. To ensure the applicability of our literature for project direction planning, we first constructed a phylogenetic tree based on molecular biological markers using the research subjects of the collected literature, aiming to determine the genetic relationships among species within the genus.
We selected the elongation factor Tu (tufA) gene—a commonly used molecular marker—and the resulting phylogenetic tree is presented below:
Fig. A phylogenetic tree based on the elongation factor Tu (tufA) gene
PQ733686.1 Ulva australis, PQ733771.1 Ulva compressa, MH538578.1 Ulva fenestrata, PQ733813.1 Ulva intestinalis,HQ610368.1 Ulva linza, PP763990.1 Ulva ohnoi, HQ610404.1 Ulva prolifera, MF172082.1 Ulva fasciata
These percentages represent bootstrap support values. A common empirical criterion is as follows: values ≥70% are considered acceptable, while those ≥90% are deemed highly reliable. The gray numbers adjacent to the branches indicate branch lengths, which correspond to the extent of sequence divergence. A value close to 0 signifies minimal divergence and a very close genetic relationship, whereas a larger value indicates greater sequence divergence.
From the phylogenetic tree, we can conclude that the genetic relationships among species within the genus Ulva are very close. Notably, Ulva linza and Ulva prolifera—the key species involved in sedimentation—form a monophyletic group.
Two novel salt tolerance genes, encoding a putative general stress protein (GspM) and an enoyl-CoA hydratase (EchM)[3], from the pond water metagenomic library are applied here.
GspM contains a complete general stress protein family GsiB[4][5] domain (COG3729) but differs from characterized GsiB (e.g., Bacillus subtilis glucose starvation-induced protein) in sequence—lacking perfect repeats, though it has three R(K/T)GG motif repeats. Phylogenetic analysis places it and its homologs in a distinct clade, separate from GsiB, YciG, or late embryogenesis abundant proteins, with origins possibly in alpha-proteobacteria from pond water.
It is chosen as a salt-tolerance gene likely due to its association with stress-responsive protein families, suggesting a role in mitigating osmotic stress, supported by its identification from a metagenome adapted to fluctuating environmental conditions.
EchM is an enoyl-CoA hydratase with 24–35% sequence identity to known E. coli enoyl-CoA hydratases (highest 35% with FadJ) and forms a distinct clade in phylogenetic analysis, indicating unique substrate specificity.
It is selected as a salt-tolerance gene because:
1) It may function in lipid/fatty acid degradation (a key process in osmotic stress adaptation by regulating membrane lipids);
2) It contains a conserved CaiD domain, suggesting potential involvement in compatible solute (e.g., L-carnitine) synthesis, which aids osmotic balance under salt stress. [6]
Fig3. The function of the salt-tolerant genes
To avoid dehydration, the osmotic pressure inside the cell increases, and certain solutes accumulate at high cytoplasmic levels through absorption from the culture medium or de novo synthesis.
The LOVdeg tag, an optogenetic protein regulatory tool, enables blue light-controlled degradation of target proteins through fusion of LOV2 domain with artificial degradation signals (e.g., EAA sequences). In darkness, LOV2's Jα helix masks these signals; 450-490 nm blue light induces helix unwinding, exposing signals for ClpXP protease degradation, enhancing efficiency 2-20-fold with high spatiotemporal precision[7]. As a blue light inducible suicide component, fusing it to tetR creates a light-controlled switch: stable in darkness to sustain cells, but blue light triggers degradation of essential proteins, causing cell death. Valuable in synthetic biology, this system can be engineered into bacteria for biosafety—natural blue light initiates suicide upon accidental environmental release.
Fig4. The principle of LOVdeg tag.
To ensure system biosafety, we designed a blue light-inducible suicide system. For light induction, particularly blue light, the pDawn promoter is commonly considered, but it operates via a three-level cascaded system with suboptimal efficiency and non-negligible leakage. Thus, we expected to seek a more efficient, less cascaded blue light induction strategy through literature research, as an innovative improvement.
We identified the LOV protein, whose Jα helix undergoes conformational unwinding under blue light. Scientists exploited this property by incorporating protease-recruiting short peptides into its conformationally variable domain, forming the LOVdeg tag. By fusing this tag to target proteins, we achieve blue light-triggered specific degradation of the target.
A repressible promoter is required to drive the suicide pathway. Among classic systems like lac and Tet operons, we selected the Tet operon to avoid interference from endogenous LacI expression in E. coli. The Tet repressor (TetR), a dimer, is tagged with LOVdeg. Under blue light, TetR bound to the operator is degraded, rendering the Tet operon light-inducible—an innovative shift from chemical induction to optogenetics.
Downstream, we employed MazF, a toxin from the toxin-antitoxin system, which degrades RNA to induce cell death.
Fig5. Overview of L.U.C.I.A. (Light Unlocks Cytotoxic Inducible Adaptor)
Choosing Ag-Nb for programmable biofilms offers key advantages:
(1) Orthogonality and composability enable precise control over adhesion specificity, allowing simultaneous function of multiple adhesin pairs;
(2) Compatibility with cell growth and division maintains adhesion during proliferation;
(3) Genetic encoding and synthetic biology standard compatibility facilitate complex multicellular design.
Fig6. Nb-Ag interactions between cells can mediate production of microscopic patterns (spatial organization of cell types, denoted in color) and morphologies (overall spatial structure of all cells, denoted by gray background).
Fig7. The adhesin pair is chose to be used to expand adhesion capabilities.
The development of a polycistronic structure of Ag43 and OmpA genes to enhance Escherichia coli biofilms, is fundamentally reliant on the combined effects of functional complementarity of the two proteins, and the benefits of a polycistronic system.
Fig8. The location and morphology of two kinds of membrane proteins.
Fig9. Ideal synergistic mechanism for biofilm growth.
Meanwhile, the polycistronic structure integrates both genes into a single vector, which avoids plasmid incompatibility issues and ensures synchronized Ag43 and OmpA expression at an optimal ratio. This strategy not only simplifies experimental procedures and reduces the metabolic burden on the host bacteria but also leverages Ag43’s autotransporter capability for autonomous surface display and OmpA’s high native compatibility with E. coli. Together, these features work synergistically to enhance biofilm formation, structural integrity, and environmental adaptability.
The settlement of Ulva zoospores involves a tightly regulated, multi-stage sequence[14][15]:
This stage marks the start of zoospore settlement, characterized by two core behaviors: first, zoospores exhibit negative phototaxis, directing themselves toward low-light regions; second, they respond to both physical surface cues (e.g., wettability, ~5 μm topographical features) and chemical/biological signals (e.g., bacterial signals, biofilms).
Upon detecting a suitable surface via apical rotational contact, zoospores undergo irreversible adhesion within 30–60 seconds. This commitment is driven by intracellular Ca²⁺ signaling, which triggers a series of coordinated physiological changes: rapid internalization of the plasma membrane, exocytosis of Golgi-derived vesicles to release glycoprotein adhesives, shedding of flagellar sheaths, retraction of flagellar axonemes, shape transition from elongated to spherical, and initiation of de novo cell wall synthesis.
The glycoprotein adhesives secreted by zoospores exhibit strong polarity: they form larger pads on hydrophilic surfaces and smaller pads on hydrophobic surfaces. This adhesive undergoes distinct time-dependent curing: it remains in a "fresh" state (with layered modulus distribution) within the first 15 minutes, forms a rigid outer "shell" after 1 hour, and shows a 65% drop in adhesion strength by 60 minutes.
Despite the early drop in adhesion strength, the zoospores’ long-term resistance to detachment improves significantly: after 8 hours of attachment, they can no longer be removed even under pressures exceeding 250 kPa. Molecular genetic evidence confirms the involvement of the hydroxyproline-rich glycoprotein (HRGP) superfamily in this process—these proteins, homologous to cell wall components in other green algae, form a physical continuum from the zoospore cell to the substrate via cross-linking, ensuring stable attachment in turbulent natural environments[16].
Fig10. The stages of Ulva spores sedimentation.
Fig11. Sketch map of spores sedimentation.
The choice of arginine[17] - rich short peptides to induce spore settlement is based on several key factors.
Fig12. The sketch map of CsgA-Arginine oligopeptide.
Fig13. The location of CsgA.
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