Structural basis of dynamic P5CS filaments

The bifunctional enzyme Δ1-pyrroline-5-carboxylate synthase (P5CS) is vital to the synthesis of proline and ornithine, playing an essential role in human health and agriculture. Pathogenic mutations in the P5CS gene (ALDH18A1) lead to neurocutaneous syndrome and skin relaxation connective tissue disease in humans, and P5CS deficiency seriously damages the ability to resist adversity in plants. We have recently found that P5CS forms cytoophidia in vivo and filaments in vitro. However, it is difficult to appreciate the function of P5CS filamentation without precise structures. Using cryo-electron microscopy, here we solve the structures of Drosophila full-length P5CS in three states at resolution from 3.1 to 4.3 Å. We observe distinct ligand-binding states and conformational changes for the GK and GPR domains, respectively. Divergent helical filaments are assembled by P5CS tetramers and stabilized by multiple interfaces. Point mutations disturbing those interfaces prevent P5CS filamentation and greatly reduce the enzymatic activity. Our findings reveal that filamentation is crucial for the coordination between the GK and GPR domains, providing a structural basis for the catalytic function of P5CS filaments.

Previous studies have revealed a characteristic compartmentation of enzymes via filamentation (Hunkeler et al., 2018;Johnson and Kollman, 2020;Liu, 2010;Park and Horton, 2019;Stoddard et al., 2020). This filamentous structure is membraneless and termed the cytoophidium for its appearance (Liu, 2010;Liu, 2016). The cytoophidium has emerged as a mechanism for the regulation of

Overall structures of P5CS filaments
The P5CS molecule contains two domains, GK and GPR, catalyzing the first and second steps in the biosynthesis of proline from glutamate. The GK domain catalyzes glutamate phosphorylation, and the GPR domain catalyzes the NADPH-dependent reduction of γ-glutamyl phosphate (G5P) to glutamate-γ-semialdehyde (GSA). The end product P5C, formed by a spontaneous cyclization reaction of GSA ( Figure 1A), will be used by another enzyme P5C reductase (P5CR) to produce proline.
In order to solve the structure of P5CS filaments, we expressed and purified Drosophila melanogaster full-length P5CS proteins. First, we analyzed the APO and substrate-bound states of P5CS by negative staining (Figure 1-figure supplement 1A-D). In our previous study, we found that Drosophila P5CS in the APO state is hard to form filaments at low concentrations (<0.05 μM). The addition of glutamate to the P5CS samples induces micron-scale filaments . Here, we observe that increasing P5CS concentration (>1 μM) also promotes the formation of filaments in the APO state. Our results show that the P5CS proteins can be self-assembled into filaments without ligands, and adding all substrates increases the length of filaments at the same concentration of the P5CS proteins. Consistent with our previous study, glutamate (a substrate of P5CS) promotes the formation and maintenance of Drosophila P5CS filaments .
Subsequently, samples of the P5CS proteins incubated with different combinations of substrates were prepared for cryo-EM (Figure 1-figure supplement 1E-J). Filaments in three conditions with (1) glutamate (P5CS Glu ), (2) glutamate and ATPγS (P5CS Glu/ATPγS ), and (3) glutamate, ATP, and NADPH (P5CS Mix ) were imaged in cryo-EM for single-particle analysis (SPA). Long and flexible filaments of P5CS were observed under all the three conditions. After 3D classification and 3D reconstruction, the electron density maps of the P5CS Glu , P5CS Glu/ATPγS , and P5CS Mix filaments reached resolutions of 4.0 Å, 4.2 Å, and 3.6 Å, respectively ( Figure 1B-D, Figure 1-figure supplements 2-4). Using a separate focused refinement strategy, we obtained multiple conformational states of the GK domain tetramer (3.1-3.5 Å) and the GPR domain dimer (3.6-4. 3Å). The cryo-EM data and model refinement statistics are provided in Table 1. The N-terminus (residues 1-44) and three disordered segments in regions I, II, and III in the GK domain were invisible in our maps.
One P5CS monomer can be roughly divided into five subdomains: (1) the glutamate-binding domain (GBD) and (2) the ATP-binding domain (ABD) at the GK domain; (3) the NADPH-binding domain (NBD), (4) the catalytic domain (CD), and (5) the oligomerization domain (OD) at the GPR domain ( Figure 1A and E). In the model, two P5CS monomers dimerize through the interaction between their GPR domains, where the β21 at the CD interacts with the β24 at the OD of the other monomer ( Figure 1F). This interaction connects two groups of hairpins and maintains the homodimer structure by a hydrogen bond network. Two P5CS dimers further assemble into a compact tetramer through the interaction at the GK domains. The P5CS tetramer serves as the building block of P5CS filaments ( Figure 1G).   Figure 1-video 2). We chose the P5CS Mix filament to display the details ( Figure 1H). In the helical P5CS filament structure, the GK tetramers serve as the core of the filament, and the GPR dimers form left-handed double helix structures around the central axis. The overall diameter of P5CS filaments in all three states is 180 Å, while the helical twist is 68° and the helical rise is 60 Å ( Figure 1H).

Structural comparison of ligand-bound GK domains
The GK domain of Drosophila P5CS is conserved with the GK protein in E. coli. Alignments of sequences and structures indicate that their secondary structures are similar as both exhibit a sandwich-like α3β8α4 topological folding (Figure 2-figure supplement 1A), which is a characteristic of the amino acid kinase (AAK) family (Marco-Marín et al., 2007;Pérez-Arellano et al., 2010;Ramón-Maiques et al., 2002).
We obtained a structure of the GK domain with the binding of glutamate in the P5CS Glu filament ( Figure 2A) and a second structure of the GK domain with G5P-Mg-ADP in the P5CS Mix filament ( Figure 2B, Figure 2-figure supplement 1B). In the P5CS Glu/ATPγS filament, the ligands could not be determined due to incomplete densities ( Figure 2-figure supplement 1C). The GK domain structure of the P5CS Glu/ATPγS filament is virtually identical to that of the P5CS Mix filament (Figure 2-figure supplement 1D). We speculate that there are two ligand-binding modes (bound with Glu-Mg-ATPγS and G5P-Mg-ADP, respectively) in the P5CS Glu/ATPγS filament. These two modes may coexist in the active sites of the GK tetramer, thereby affecting the 3D reconstruction of the structures. The unexpected presence of G5P could be due to the contamination of ATP in the commercial ATPγS (80% pure) and all substrates were in excess during our sample preparation. Thus, no ligand was modeled in the GK domain structure of the P5CS Glu/ATPγS filament.
In the GK domain, a valley-like pocket locates between GBD and ABD, providing the binding sites for glutamate, ATP, or their derivatives ( Figure 2C and D). Glutamate binds to the active site of GBD vertically (Figure 2A and C, Figure 2-figure supplement 1E). In contrast, G5P and ADP extend towards each other in the P5CS Mix filament, and glutamate at the binding site is converted into the intermediate G5P. At ABD of the P5CS Mix filament, the phosphate donor ATP becomes an ADP, associating with an Mg 2+ ( Figure 2B and D).
Superimposing the GK tetramer in the P5CS Glu filament and that in the P5CS Mix filament revealed that the major motion of the GK domain occurred at the region containing flexible loops or disordered segments, whereas the α3β8α4 fold showed minor movement ( Figure 3A). Meanwhile, based on the disorder densities in region II (Figure 3-figure supplement 1A-D), we modeled the possible trend of the missing segment with a dashed line ( Figure 3A). In the P5CS Glu filament, we speculate that the disordered segment in region II acts as a closed loop, which traps glutamate in GBD (Figures 2C and 3A). In the P5CS Mix filament, the same segment shifts away from the top of the binding pocket and domain (OD) of the GPR domain are shown in cyan, purple, and pink, respectively. Bifunctional P5CS enzyme catalytic reaction and residue numbers for domain boundaries are shown. (B-D) Single-particle analysis for 3D reconstruction of P5CS filaments, three cryo-EM maps of P5CS Glu filament, P5CS Glu/ ATPγS filament, and P5CS Mix filament are colored by local resolution estimations. (E) The structures of the P5CS monomer and color codes for P5CS models are indicated. (F) The P5CS dimer. Two monomers (gray or color coded by domain) interact via GPR domain hairpins contact. (G) The P5CS tetramer (sphere representation) is formed via GK domain interaction (cartoon representation) between two P5CS dimers (gray or color coded by domain). (H) The sphere and cartoon representation of P5CS filaments. P5CS filaments are modeled by the cryo-EM map. The rotated view is shown in the right panel; its rise, twist, and width are indicated.
The online version of this article includes the following video and figure supplement(s) for figure 1:       forms an open loop, in which residue M213 interacts with G5P ( Figure 3A, Figure 3-figure supplement 1E). We notice that the closed loop has a steric clash with G5P, preventing the binding of G5P under such a conformation ( Figure 3-figure supplement 1D). Our findings support the idea that region II at the GK domain engages in regulating the catalytic reaction.   The helix-helix structure (residues 105-113, 115-124) at region I of the P5CS Glu filament transforms into a helix-loop-helix structure (residues 105-119, 120-122, 123-128) in the P5CS Mix filament ( Figure 3A, Figure 3-figure supplement 1F). This helix-loop-helix structure is referred to as the 'hook' structure. The transformation of the hook structure results in new contact sites between neighbor tetramers in the vertical direction, which is evidenced by a rigid density in our map ( Figure 1B-D, Figure 1-figure supplements 2 and 3).
On the other hand, we notice the conformational variation of the loop at region III and a loop (residues 64-77) of GBD shifting greatly by approximately 3 Å away ( Figure 3A). The function of these conformational changes is unclear, which may relate to conformational changes of the active site. In order to investigate the conformational changes involved in the catalytic reaction, we further compared the tetramer structures of the GK domain in the P5CS Glu and P5CS Mix filaments ( Figure 3B). The GK domain of each protomer rotates approximately 10° around its central axis, causing the horizontal compression of the GK domain dimer. By comparing the structures of the GK domain with various ligands, we demonstrate the conformational changes, which may be associated with phosphorylation of the substrate glutamate.

Open and closed conformations of GPR domains
The GPR domain of P5CS belongs to the aldehyde dehydrogenase (ALDH) superfamily. ALDH family uses NAD(P) + to catalyze the conversion of various aldehydes into their corresponding carboxylic acids. Many studies on ALDHs have shown that a conserved residue cysteine acts as the active site of nucleophile, forming thiohemiacetal intermediate with substrate (Koppaka et al., 2012;Liu et al., 1997;Perozich et al., 1999). Curiously, the NADPH-utilizing GPR domain of P5CS catalyzes the reverse reaction of ALDHs.
On the basis of P5CS structures, we display four different binding modes ( Figure 4A-D) of the GPR domain. In the P5CS Glu filament, no ligand binds to the GPR domain ( Figure 4A). In the P5CS Glu/ ATPγS , however, we observed the density of a G5P at the CD active site ( Figure 4B, Figure 4-figure supplement 1A). It might be a contamination of ATP, leading to the production of the substrate G5P. In this model, the binding mode of G5P (referred to as the G5P-binding state) is clearly solved. By focus refinement of the GPR dimer in the P5CS Mix filament, we determined two additional states of the GPR domain ( Figure 4C and D). One is the NADP(H)-binding state, when NADP(H) is present at NBD (Figure 4-figure supplement 1B). The other is the NADP(H)-released state, of which the cofactor binding site is empty.
Conformational comparison of unliganded, G5P-binding and NADP(H)-released states shows that the overall structures of the GPR domain are similar (Figure 4-figure supplement 2A). The structure      https://elifesciences.org/articles/76107/figures#fig4video1 of the NADP(H)-release state, which has no bound ligand, is identical to the unliganded state. The binding of G5P to CD of the GPR domain does not lead to obvious conformational changes.
Next, we compared the structures of the G5P-binding state and NADPH-binding state ( Figure 4E, Figure 4-figure supplement 2B). We have found that the structures of CD and OD are generally consistent in those two states, while NBD of the GPR domain in those two states differs greatly ( Figure 4E). NBD contains consecutive alternating α-helices and β-strands (α 2 -β 5 -α 2 ) architecture, which is known as the Rossmann-like fold for dinucleotide binding (Cheek et al., 2005). By superimposing the Rossmann-like fold and the entire NBD, we determined conformational changes between the GPR domain at the G5P-binding state and that at the NADP(H)-binding state ( Figure 4F).
Upon NADP(H) binding, the residue R525 interacts with the adenine moiety. This interaction transforms the 525 REE 527 loop into an ordered structure that extends the α16 helix in the Rossmann-like fold ( Figure 4F). Meanwhile, the entire NBD rotates approximately 15° along the cylinder axis (Figure 4figure supplement 3A) and slides towards CD ( Figure 4F; Figure 4-video 1). We hypothesize that the helix, when turns disordered, loses contact with the adenine moiety and then separates the cofactor from NBD via a conformational selection mechanism. A similar phenomenon was also observed in ALDH1L1 (Tsybovsky and Krupenko, 2011). This transformation contributes to bringing the nicotinamide ring of the NADP(H) close to the catalytic residue C598 of CD. Conformational changes triggered by the binding of NADPH subsequently initiate the transfer of the hydride ion from NADPH to the intermediate G5P (Figure 4-figure supplement 3B).      In our models, unliganded, G5P-binding and NADP(H)-released states represent as open conformation, and the NADP(H)-binding state represents as closed conformation. We propose that the P5CS filament accommodates the GPR domain in both open and closed conformations. Therefore, recurring transformations between these two conformations are essential for the catalytic cycle at the GPR domain.

Filamentation regulates the enzymatic reaction
In P5CS Glu , P5CS Glu/ATPγS , and P5CS Mix filaments, neighboring GPR dimers interact with each other and form a helical structure. The interaction formed between F642-P644 of the contact loops in adjacent GPR domains, which appears as a CH/Pi interaction (Zondlo, 2013), is critical for the filamentation ( Figure 5A and B, Figure 5-figure supplement 1). In P5CS Glu/ATPγS and P5CS Mix filaments, the additional interface between hook structures pairs locks adjacent GK tetramers. The hook structure extrudes from the GK domain. In a GK tetramer, four hooks extrude toward two opposite directions to form a 'spinning top' arrangement. Therefore, two pairs of hooks in a GK tetramer interact with their counterparts in two adjacent GK tetramers ( Figure 5A and C). The hook interaction forms strong contacts via hydrogen bonds (M119-R124, L121-M123) and salt bridges (E116-R124). Therefore, a combination of the GPR contact (for the double helix) and GK contact (for the axis) stabilizes the P5CS filament.
To understand the function of P5CS filaments, we generated two mutants, R124A and F642A, which are predicted to abrogate the tetramer-tetramer contact sites of the GK domains and GPR domains, respectively. Negative stain of the mutant P5CS showed that the P5CS F642A mutant proteins did not assemble into a filament with or without ligands ( Figure 5-figure supplement 2A). These results indicate that the interaction at the GPR domain interface is crucial for P5CS filamentation.
In contrast, the P5CS R124A mutant proteins formed long filaments in the APO state as well as in the presence glutamate ( Figure 5-figure supplement 2B). We observed that glutamate-bound P5CS R124A filaments disassembled at the initial phase of adding ATP. Being incubated with all substrates, P5CS R124A formed shorter filaments than P5CS WT ( Figure 5-figure supplement 2B).
We propose that the interactions among the hook pairs are required for stabilizing the filament during the transformation from the P5CS Glu filament to the P5CS Glu/ ATPγS filament or P5CS Mix filament. We subsequently analyzed the activity of the wild-type P5CS and two mutants, R124A and F642A. The two mutants exhibited a dramatically compromised activity in comparison with the wild-type P5CS ( Figure 5D), suggesting that the integrity of filament is essential to the catalytic reactions.

The GK domain
We observed two ligand-binding modes in the GK domain. Due to the lack of ATP-bound structure, it is difficult to determine whether ATP plays a decisive role in these conformational changes. According to a previous report on the N-acetyll-glutamate kinase (NAGK), nucleoside is important for the conformational change of the AAK domain, and the structures are similar when bound by ADP or AMPPNP (Gil-Ortiz et al., 2011). Based on the similarity of sequences and structures between GK and NAGK (Marco-Marín et al., 2007), we propose that the conformation of the GK domain in the P5CS Glu filament would transform upon the binding of ATP, thereby triggering the formation of hook structure and completing the catalytic reaction. Although we solved the clear structure of the P5CS Glu filament, further research is needed to understand how the conformation of glutamate binding contributes to the extension of P5CS filaments.

The GPR domain
Aspartate-β-semialdehyde dehydrogenase (ASADH) catalyzes NADPH-dependent reductive dephosphorylation of β-aspartyl phosphate to aspartate-β-semialdehyde (Karsten and Viola, 1991). The GPR domain of P5CS and ASADH catalyzes the same type of reaction. Interestingly, the binding of NADP + will change the cofactor binding domain of ASADH from open conformation to closed conformation (Hadfield et al., 2001). Thus, we speculate that their catalytic mechanisms have something in common. In the GPR domain of Drosophila P5CS, our data suggest that the catalytic residue C598 of CD attacks the G5P to form the first tetrahedral thioacetal intermediate in the reaction, and then expulsion of phosphate collapses to form a stable thioacyl enzyme intermediate. A hydride is then transferred to this intermediate from NADPH, with subsequent collapse to release the product GSA. Furthermore, the NADPH-binding site is located inside the filament, close to the GK domain. The G5P binding site is close to the external solution environment, which is proposed to facilitate the release of the product GSA/P5C ( Figure 5-figure supplement 3, Figure 5-video 1). G5P can freely bind to the GPR domain in our model (G5P-binding state). However, in the closed conformation, when the nicotinamide ring of NADP(H) approaches the G5P-binding site, the substrate tunnel entrance may be blocked by NADP(H). This may affect the subsequent binding of G5P. Therefore, we speculate that the GPR domain should bind with G5P prior to NADPH binding. However, whether this mechanism is a preferred binding order needs to be further verified by kinetic experiments.

The P5CS filament
As mentioned in the 'Results' section, we observed that mutated residues R124A and F642A do not directly participate in the active sites, while they are crucial for filamentation. This suggests that the P5CS filamentation couples the reaction catalyzed between the GK domain and GPR domain through transferring unstable intermediate G5P (Pérez-Arellano et al., 2010;Seddon et al., 1989). Considering the distance between the GK and GPR domains is about 60 Å ( Figure 5-figure supplement 3, Figure 5-video 1), we propose a model that P5CS filament may exhibit a scaffold architecture that stabilizes the relative position of the GK and GPR domains, the cooperation between which may produce electrostatic substrate channels that mediate the transfer of unstable intermediate G5P.
In addition, P5CS filamentation may create a half-opened chamber with the active sites located at the inner part of the filament. Since the GK domain is catalytically faster than the GPR domain, the unstable intermediates G5P accumulate within the filament. This microenvironment may reduce the amount of G5P escaped into the solvent, thereby facilitating the rate-limiting reaction at the GPR domain.

The working model
Together, we propose a coupling catalytic reaction mechanism of Drosophila dynamic P5CS filament. In this proposed model, spontaneous filamentation occurs at the APO state, and elongation of P5CS filament is associated with the binding of glutamate. Upon the binding of glutamate, the binding pocket at the GK domain is bound by ATP; subsequently, conformational changes facilitate the formation of a hook structure and phosphorylation of glutamate, which produces G5P. When products of the GK domain dissociate from the active site, G5P would be trapped by the channel and chamber within the filament and further captured by the GPR domain. Next, NADPH binds to the GPR domain, triggering the conformational change into closed conformation, which brings the NADPH towards the catalytic residue C598 and facilitates the reaction. After this reaction, NADP + and GSA will be released, and the GPR domain returns to its open conformation ( Figure 6). This working model suggests that the GK and GPR domains undergo continuous conformational transition during catalysis, resulting in a dynamic filament.
In the P5CS Glu filament, the GK and GPR domains are likely in a stable conformational state, while vibration may occur in the GPR domain of P5CS Glu/ATPγS and P5CS Mix filaments due to the binding of ligands. This notion could be supported by the differences in their local resolution ( Figure 1B-D). We speculate that the swing of GPR in the catalytic reaction could destabilize the interaction between adjacent GPR domain dimers in the filament. Therefore, the extra interaction at the hook structure of the GK domain may be required for the stabilization of the filament. This proposed stabilization is consistent with negative stain data showing that the P5CS R124A mutant cannot stabilize the filament structure in the catalytic process and lose the ability to form the long filaments.

P5CS and human disease
Recently, accumulative evidences have shown that mutations on the human P5CS gene (ALDH18A1) is one of the causes of hyperammonemia, neurocutaneous syndromes, and motor neuron syndrome (Baumgartner et al., 2005;Magini et al., 2019;Marco-Marín et al., 2020;Pérez-Arellano et al., 2010). Such mutations may result in the loss of P5CS function in various degrees. Drosophila P5CS residue R124 in region I of the GK domain, which corresponds to R138 of human P5CS, is highly conserved among different eukaryotes ( Figure 5-figure supplement 4). However, 17 residues in region I, including the hook structure, are absent in E. coli GK. Pathogenic mutations of R138 in human P5CS, which are proven to be the cause of autosomal-dominant cutis laxa, have been demonstrated with a decreased activity and a dispersed distribution in mitochondria (Fischer-Zirnsak et al., 2015;Yang et al., 2021). In the protein structure database, there is only the GPR domain structure available for human P5CS (PDB: 2H5G). Its overall structure is similar to the GPR domain of Drosophila P5CS (Figure 4-figure supplement 2B). Although it is still unknown whether human P5CS can form filament structure in vitro, it is reasonable to suspect that the filament-forming property is conserved between human and Drosophila P5CS based on their structural similarity. Our structure reveals that the R138 mutation on human P5CS could abrogate the interaction between hook structures of GK domains, and thereby destabilize the filament and coupling of reactions at the two domains.
In summary, our cryo-EM structures of Drosophila P5CS filament present the assembly mode of P5CS protein and provide a molecular basis for a further understanding of the reaction mechanism of the GK and GPR domains. In our proposed model, the coupling of the GK and GPR domains in the filamentous structure facilitates the catalytic reaction of the bifunctional enzyme P5CS. Additional structural studies of P5CS filaments are required to determine whether there is an underlying regulatory mechanism that transmits information between the GK and GPR domains in the tetramer and along the filament.

P5CS protein purification
The full-length D. melanogaster P5CS gene was cloned into a modified pET28a vector with a 6 × His SUMO tag fused at the N terminus; the fusion proteins were expressed in E. coli Transetta (DE3) cells overnight at 16°C after induction with 0.1 mM IPTG at OD 600 range of 0.6-0.8. The remainder of purification was performed at 4°C. The harvested cells were sonicated under ice and purified by Ni-NTA agarose beads (QIAGEN) in lysis buffer (50 mM Tris-HCl pH 8.0, 500 mM NaCl, 10% glycerol, 20 mM imidazole, 1 mM PMSF, 5 mM β-mercaptoethanol, 5 mM benzamidine, 2 μg/ml leupeptin, and 2 μg/ml pepstatin). After in-column washing with lysis buffer, the proteins were eluted with elution buffer (50 mM Tris-HCl pH 8.0, 500 mM NaCl, 250 mM imidazole, 5 mM β-mercaptoethanol), peak fractions were treated with SUMO protease for 1 hr at 8°C. The P5CS proteins were further purified through HiLoad 16/600 Superdex 200pg gel-filtration chromatography (GE Healthcare) in column buffer (25 mM HEPES pH 7.5 and 100 mM KCl), peak fractions were collected, concentrated, and stored at -80°C before use.

Enzyme activity assays
The full-length wild-type or mutant P5CS (100 nM protein) activity was determined in the reaction buffer containing 25 mM HEPES pH 7.5, and 10 mM l-glutamate (Sigma), with added 20 mM MgCl 2 , 10 mM ATP (Takara), and 0.5 mM NADPH (Roche) used to initiate the reaction (Magini et al., 2019;Sabbioni et al., 2021), then the reaction was monitored at 37°C in an MD-SpectraMax i3 plate reader and absorbance at 340 nm was measured every 20 s for 10 min (one experiment, n = 3). The NADPH concentration was converted from A340 with the standard curve determined at the same experiment.

Negative staining
Wild-type or mutation P5CS proteins were mixed with different substrate conditions. In brief, the final concentration was as follows: 25 mM HEPES pH 7.5, 100 mM KCl, 10 mM MgSO 4 , 100 mM l-glutamate, 10 mM ATP, and 0.5 mM NADPH. The prepared protein samples were applied to glow-discharged carbon-coated EM grids (400 mech, EMCN), and stained with 1% uranyl acetate. Negative-stain EM grids were photographed on a Tecnai Spirit G21 microscope (FEI).

Cryo-EM grid preparation and data collection
For cryo-EM, purified full-length P5CS was diluted to approximately 2 μM and dissolved in buffer containing 25 mM HEPES pH 7.5, 100 mM KCl, 10 mM MgSO 4 , and incubated with 20 mM l-glutamate for the P5CS Glu filament preparation. The P5CS Glu/ATPγS filament was added with an additional 0.5 mM ATPγS (Abcam) compared to the P5CS Glu filament. For the P5CS Mix filament, P5CS proteins (2 μM) were incubated with 100 mM KCl, 10 mM MgSO 4 , 20 mM l-glutamate, 2 mM ATP, and 0.5 mM NADPH. All the samples were incubated for 1 hr on ice before vitrification. The P5CS filament samples were placed on H 2 /O 2 glow-discharged holey carbon grids (Quantifoil Cu 300 mesh, R1.2/1.3) or amorphous alloy film (CryoMatrix M024-Au300-R12/13). Then, the grids were immediately blotted for 3.0 s and plunge-frozen in liquid ethane cooled by liquid nitrogen using Vitrobot (Thermo Fisher) at 4°C with 100% humidity. Images were collected on Titan Krios G3 (FEI) equipped with a K3 Summit direct electron detector (Gatan), operating in counting super-resolution mode at 300 kV with a total dose of 72 e − /Å 2 , subdivided into 50 frames in 4 s exposure using SerialEM (Mastronarde, 2005). The images were recorded at a nominal magnification of 22,500 × and a calibrated pixel size of 1.06 Å, with defocus ranging from 0.8 to 2.5 μm.

Image processing and 3D reconstruction
The whole-image analysis was performed with RELION3 (Zivanov et al., 2018). We used MotionCor2 (Zheng et al., 2017) and CTFFIND4 (Rohou and Grigorieff, 2015) via RELION GUI to pr-process the image, movie frames were aligned, and the contrast transfer function (CTF) parameters were estimated in this process. After manual selection, there are 4933 images for the P5CS Glu dataset, 6408 images for the P5CS Glu/ATPγS dataset, and 10,566 images for the P5CS Mix dataset left for further processing. For the flexibility of P5CS filaments, SPA was carried out in our reconstructions and no helical symmetry was implied in the whole process. Reference-free particle picking built in RELION3 was performed. This process provides 1,994,786 particles for P5CS Glu , 2,024,372 particles for P5CS Glu/ATPγS , and 8,027,582 particles for P5CS Mix . At first, the particles were extracted binning two or three times for the fast 2D classifications. Datasets were cleaned with several rounds of 2D classification and the bin factors were gradually reduced to one at the same time. After 3D classifications with C1 symmetry were applied, several classes were selected to do finer 3D classifications with D2 symmetry. Classes with the intact structure were retained for 3D refinement with D2 symmetry. For the 3D refinement, 432,746, 327,841, and 1,412,498 particles were used for each dataset. The maps including three P5CS tetramer layers were obtained. The relative motion between GK and GPR limited the refinement at a high resolution, so we used the partition reconstruction strategy to improve the resolution for both the GK and GPR domains. For the GK domain, we used continued local refinement to improve the resolution with a mask focus on the middle layer GK. Then, the Ctf-refinement and Bayesian polishing were performed for the remained particles and improved the resolution to 4. 1Å, 4.1 Å, and 3.6 Å for three-layer P5CS maps and 3.5 Å, 3.4 Å, and 3.1 Å for GK maps. For the GPR domain, particles were expended symmetry for the 3D classification without alignment. Several classes with the intact structure were selected and oriented; symmetry collapse was done at the same time. Then, 3D classifications and refinements with C2 symmetry were performed. For the P5CS Mix , two different states of GPR were captured. Finally, we got 286,291, 348,804, 193,482, and 233,624 particles to construct maps for the GPR domain with 3.6 Å, 4.2 Å, 4.3 Å, and 4.2 Å resolutions. LocalRes was used to estimate the local resolution of our map.

Model building refinement and validation
Based on our maps with the near-atomic resolution, the model of the GK and GPR domains was generated with focused refinement maps in different states. The initial model of the GK and GPR domains was generated via swiss model regarding 4Q1T (GK from Burkholderia thailandensis) and 2H5G (human GPR domain) as a reference, respectively. Manual adjustment and building the missing regions were done in Coot (Emsley and Cowtan, 2004). Real space refinements were performed with Phenix (Adams et al., 2011). The full-length P5CS models were linked using the corresponding GK and GPR structures; the linker was generated in the Coot and refined via Phenix. Figures and movies were generated with UCSF Chimera (Pettersen et al., 2004) and ChimeraX (Goddard et al., 2018).


Physiological symmetry of transcranial magnetic stimulation‐evoked EEG spectral features

Abstract Transcranial magnetic stimulation (TMS)‐evoked EEG potentials (TEPs) have been used to study the excitability of different cortical areas (CAs) in humans. Characterising the interhemispheric symmetry of TMS‐EEG may provide further understanding of structure–function association in physiological and pathological conditions. We hypothesise that, in keeping with the underlying cytoarchitectonics, TEPs in contralateral homologous CAs share similar, symmetric spectral features, whilst ipsilateral TEPs from different CAs diverge in their waveshape and frequency content. We performed single‐pulse (<1 Hz) navigated monophasic TMS, combined with high‐density EEG with active electrodes, in 10 healthy participants. We targeted two bilateral CAs: premotor and motor. We compared frequency power bands, computed Pearson correlation coefficient (R) and Correlated Component Analysis (CorrCA) to detect divergences, as well as common components across TEPs. The main frequency of TEPs was faster in premotor than in motor CAs (p < .05) across all participants. Frequencies were not different between contralateral homologous CAs, whilst, despite closer proximity, there was a significant difference between ipsilateral premotor and motor CAs (p > .5), with frequency decreasing from anterior to posterior CAs. Correlation was high between contralateral homologous CAs and low between ipsilateral CAs. When applying CorrCA, specific components were shared by contralateral homologous TEPs. We show physiological symmetry of TEP spectral features between contralateral homologous CAs, whilst ipsilateral premotor and motor TEPs differ despite lower geometrical distance. Our findings support the role of TEPs as biomarker of local cortical properties and provide a first reference dataset for TMS‐EEG studies in asymmetric brain disorders.

| INTRODUCTION
Transcranial magnetic stimulation (TMS) coupled with EEG (TMS-EEG) is a controlled perturbational approach that directly activates a target cortical area (CA) to assess its excitability (Ilmoniemi et al., 1997;Rosanova et al., 2009). TMS-evoked EEG potentials (TEPs) are complex waveforms generated by averaging segments of EEG recording ("trials") which are time-locked to the TMS pulses. Typically, TEPs have a duration of hundreds of milliseconds and are characterised by sustained increases of power in specific frequency bands that depend on the CA targeted (Rosanova et al., 2009). When not confounded by scalp muscle artefacts, auditory, or extraneous sensory activations, TEPs provide a reliable readout of the reactivity of cortical circuits (Hallett, 2007;Komssi & Kähkönen, 2006;Rogasch & Fitzgerald, 2013;Rosanova et al., 2012). TMS can be delivered with different pulse configurations, whereby current is either delivered with a bidirectional flow (biphasic stimulation), or dampened after the first quarter cycle, thus delivering a unidirectional current flow (monophasic stimulation) (Sakai et al., 1997).
In TMS-EEG experiments, either passive or active EEG systems configurations may be used (Mancuso et al., 2021). Active configurations are more sensitive to artefacts due to transient voltage changes; however, signal pre-amplification directly at the electrode level may allow for better signal quality at higher interelectrode impedance compared to passive systems, and enables faster preparation in the clinical environment (Laszlo et al., 2014).
Previous studies employing biphasic TMS and passive electrodes have shown that TEPs follow a rostro-caudal gradient in their main oscillatory frequency (i.e., natural frequencies) (Rosanova et al., 2009). This intrinsic frequency gradient at which CAs oscillate is thought to reflect the cytoarchitectonics, as well as the connectivity, of distinct thalamocortical modules (Ferrarelli et al., 2012;Rosanova et al., 2009) and can be altered in brain pathological conditions (Ferrarelli et al., 2012). Up-to-date, this gradient has only been described using biphasic TMS and passive electrodes. Furthermore, the interhemispheric symmetry of the natural frequency gradient has never been assessed.
In this work, we sought to determine the symmetry of TEP spectral features by examining the presence of comparable bilateral natural frequencies specific to each CA, to corroborate previous work investigating the similarity of TEPs recorded from homologous regions (Casula et al., 2020;Vallesi et al., 2021). We also assessed the use of monophasic TMS coupled with active high-density EEG (hd-EEG) to reproduce the natural frequencies previously determined in premotor areas, and for the first time measured the natural frequencies for the hand region of the primary motor cortex on the left and right hemispheres. Additionally, we assessed the interhemispheric dynamics by applying the interhemispheric signal propagation (ISP) and balance (IHB) as described elsewhere (Casula et al., 2020). We extracted the spectral features of TEPs recorded from 10 healthy individuals in whom we targeted both premotor and motor CAs, and compared TEP natural frequencies and voltages by applying time-frequency analysis and Pearson's correlation coefficient (R), respectively. We also identified the TEP components that were reliably reproducible across participants and specific to the targeted CA by applying the correlated component analysis (CorrCA). Our ultimate goal was to demonstrate that TMS may serve as a potential biomarker to assess cortical activity providing functional information on the underlying cytoarchitectonics, beyond the information provided by standard neuroimaging.

| Participants
The study was approved by the local ethics committee (REC ref 15/LO/1642). All participants gave written informed consent. Exclusion criteria were as follows: history of traumatic brain injury, neurological or psychiatric diseases, presence of intracranial metallic implants, drug release dispensers, metallic tattoos or cardiac pacemakers (Rossi et al., 2009). Ten right-handed healthy participants were recruited to participate in the study (six females; age: median 33.00 years, interquartile range [IQR] 5.00, all with higher education level, Table 1).

| Experimental setup
Experiments were conducted at the Chalfont Centre for Epilepsy, Buckinghamshire, UK, under the research governance of University College London (UCL). We devised a systematic approach for our TMS-EEG assessment (Figure 1, full details of the protocol are available in the Supplementary Material S1).

| Transcranial magnetic stimulation-click sound masking
To avoid the contamination of TEPs by auditory responses to the click produced by the TMS coil, we adopted the TMS-click sound masking toolbox TAAC  and participants wore noisecancelling in-ear headphones (Shure SE215-CL-E Sound Isolating, for the detailed methodology see Supplementary Material S2).

| High-density EEG recording
For all the experimental sessions, we used hd-EEG recording following international standards (Nuwer et al., 1998;Sinha et al., 2016). An ActiCHamp 63-channel amplifier was used, and TMS-compatible acti-CAP active electrodes (Brain Products GmbH, Germany) were placed in the international 10-20 montage referenced to the forehead. The impedance of all electrodes was kept below 5 kΩ (Ilmoniemi & Kiči c, 2010). EEG signals were band-pass filtered between 0.1 and 500 Hz and sampled at 5 kHz with 32-bit resolution. With this specific equipment, the magnetic artefact induced by the stimulation pulse does not last longer than 5 ms in the EEG recording. At the start of each experiment, the motor "hotspot" for the first dorsal interosseus (FDI) muscle was located over the hemisphere contralateral to the muscle being monitored and the resting motor threshold (RMT) was determined for both hemispheres, following Rossini and colleagues' guidelines (Rossini et al., 2015). RMT was considered as the TMS intensity required to elicit motor evoked potentials (MEPs) >50 μV in 5 of 10 trials delivering pulses at 0.2 Hz minimum, measured from the FDI muscle (Supplementary Material S3).

| Brain navigation, TMS targeting and brain stimulation
In this work, CA refers to any of the motor or premotor brain regions, whilst we define the region of interest (ROI) as the group of four channels related to each CA under study (F1-Fz-Fc1-Fcz for left premotor, F2-Fz-Fc2-Fcz for right premotor, C5-C3-Cp5-Cp3 for left motor, C4-C6-Cp4-Cp6 for right motor, Table S1). Two bilateral scalp targets were identified within the areas indicated by the four abovementioned ROIs. Individual structural magnetic resonance images (MRI, T1-3D sequence) were available for five participants (Table 1).
During the assessments on these subjects, we verified that the two anterior targets were overlapping the caudal frontal superior gyrus (i.e., the premotor-BA6) and that the two posterior targets were overlapping the precentral gyrus (i.e., the motor-BA4) areas. This selection of CAs was based on previous studies that assessed cortical excitability in vivo in humans (Ferrarelli et al., 2012). In these subjects,  Table S1 for individual intensities).
A real-time visualisation tool (rt-TEP) was used to guide coil orientation and to determine the stimulator intensity, minimising muscle artefacts and ensuring the presence of a detectable TEP . Stimulation intensity was determined differently depending on the CA under study. For TEPs evoked in the motor CA, intensity was the highest available below ≤90% of the RMT, to avoid possible sensory-feedback contamination, as previously described (Fecchio et al., 2017). For premotor TEPs, we used the stimulator intensity able to elicit at least a 10 μV-amplitude response in the average of the first 20 trials. This response was measured in the channels closest to the stimulation site (as per Casarotto et al., 2022), on the first peak-to-peak component between 10 and 50 ms after the TMS pulse, using an average reference montage. In this work, we also measured the first TEP component after pre-processing and averaging. We consistently found intra-session asymmetric amplitudes with the highest voltages in the channels under the coil, which we considered as a proof of direct cortical perturbation (Belardinelli et al., 2019;Casarotto et al., 2022). Wilcoxon matched pairs signed rank test was

| Data pre-processing
TMS-EEG data pre-processing was performed using Matlab R2016a (The MathWorks). First, the TMS artefact was removed from all the trials replacing the recording between À2 and 5 ms from the TMS pulse with the 7 ms of time interval before (between À9 and À2 ms from the pulses). All trials were segmented ±800 ms around the stimulus and high-pass filtered at 1 Hz (Makeig et al., 2002). Channels containing line-noise >50 μV lasting more than the 10% of the entire session duration were manually excluded. Subsequently, segments containing more than 50 μV of electrical activity were detected and rejected by visual inspection by two trained researchers (SDA, DJJ).
Channels were re-referenced to the average reference and rejected channels were interpolated using the EEGLAB spherical interpolation function. We used independent component analysis (ICA) to remove any residual artefacts caused by eye movements, scalp muscle activation or electrical interference of devices (EEGLAB runica function; Makeig et al., 2002). Finally, data were down-sampled at 1 kHz, lowpass filtered (45 Hz, notch 50 Hz) and segmented again in a time window of ±600 ms around TMS pulses (Fecchio et al., 2017;Makeig, 1993). After pre-processing, we included 10 participants with bilateral TEPs obtained from premotor and motor stimulations for further analysis (Table 1).

| Natural frequencies
We considered the natural frequency as the main (i.e., the most powerful) frequency of TMS-evoked oscillation globally across all channels, between 20 and 200 ms after the TMS pulse (Rosanova et al., 2009).
We assessed TEP spectral features by analysing the event-related spectral perturbation (ERSP). ERSP measures the average dynamic change in amplitude across all the bands of the EEG frequency spectrum in relation to a specific event (Makeig, 1993). We performed time-frequency decomposition analysis with wavelet transformation (Morlet, 3.5 cycles) between 8 and 45 Hz by computing the EEGLAB Newtimef function (Grandchamp & Delorme, 2011), within the Matlab-based public license toolbox EEGLAB (Delorme & Makeig, 2004). Absolute spectral normalisation was conducted on all trials by performing a full-epoch length correction. A pre-stimulus baseline correction between À500 and À100 ms from the pulse was applied to the pre-processed data, with the time window set 100 ms apart from the stimulus in order to avoid any possible post-stimulus contamination. The Newtimef function computes the surrogate distribution (i.e., creates timepoints with values randomly chosen from the baseline) at each frequency by permuting real baseline values.
Thus, it tests whether the original ERSP value points are present in the 99.5% tail of the surrogate distribution of any given frequency (Fecchio et al., 2017), in which case each specific time-frequency point is considered significant at α < .01 (Fecchio et al., 2017), after correction for multiple comparisons, and using the false discovery rate (FDR) procedure (Grandchamp & Delorme, 2011

| Correlation coefficient analysis
We measured the degree of correlation between the voltages of the Since we adopted two different approaches for tuning the stimulation intensity across different areas, we controlled for the possible association between the natural frequencies and the stimulation intensities used. To this aim, we calculated the correlation coefficient between the absolute values of natural frequencies and intensities used in each area under stimulation. We then conducted a linear regression analysis of the distribution (y = α + βx, where y is intensity and x is the natural frequency) to further analyse the association between intensity stimulation and natural frequencies. Values of p < .05 were deemed as significant.

| Correlated component analysis
We

| Interhemispheric signal propagation and inter hemispheric balance
To assess the interhemispheric transmission, we first analysed the TMSevoked response on the stimulated hemisphere and on the contralateral.  Figure 2). We obtained similar results when comparing natural frequencies calculated at the local level (full results available in the Supplementary Material S6, Figure S1, Table S2). Additionally, we found that variability in the natural frequencies was not affected by the use of individual MRI versus template (details in Table S3).

| Correlation coefficient analysis
The Pearson's coefficient (R) analysis showed higher correlation between contralateral homologous ROI comparisons ( Figure 3)

| Amplitude of the first TEP component after pre-processing and averaging
For this component, we found larger amplitude in premotor TEPs than There was a significant difference between left ipsilateral premotor and motor TEPs (W = À39.00, p = .0488) and between right ipsilateral premotor and motor TEPs (W = À53.00, p = .0039).

| Local mean field power
In order to control for the potential confound of the stimulation impact on the reactivity of each stimulated CA, we calculated and compared the local mean field power (LMFP) of each TEP (detailed methodology and results in Supplementary Material S9, Figure S3).

| Interhemispheric signal propagation and interhemispheric balance
We calculated the ISP from the left and right motor cortices. We found a consistent interhemispheric inhibition across all our participants (see Supplementary Material S11).

| TEP differentiation and symmetry
Here, we used single-pulse (<1 Hz) navigated monophasic TMS to stimulate bilateral cortical areas and simultaneously record hd-EEG with active electrodes. Using a different TMS pulse configuration and a different amplifier to those in previous reports, we first reproduced previous findings showing the presence of distinct frequencies evoked across different CAs perturbed by direct transcranial magnetic pulses (Sakai et al., 1997;Sommer et al., 2006). This result confirms the existence of a specific frequency tuning, called the natural frequency, that is intrinsic to the targeted circuit and independent of the specific stimulation and recording set-up (Ferrarelli et al., 2012;Rosanova et al., 2009). Most importantly, we find that such tuning is symmetric across the two hemispheres.
LMFP analysis showed no significant difference in the response to TMS between 20 and 200 ms across the different stimulation targets. Therefore, the natural frequencies observed are unlikely to reflect differences in the overall effectiveness of the TMS perturbation. Our results showed symmetrical natural frequencies and a high correlation coefficient between contralateral homologous ROIs (Figures 2 and 3). We found significant differences when comparing the distance between homologous contralateral motor areas (which was higher) and the other distances ( Figure 5). However, motor TEPs showed symmetrical spectral frequencies despite higher geometrical distance between the contralateral homologous motor targets, whilst ipsilateral motor and premotor TEPs were different despite their closer proximity. These findings confirm that differences in the TEPs features depend more on the cytoarchitectonic differences of the underlying cortex rather than on the geometrical distance between the stimulated areas. Cytoarchitecture and histology are known to be similar in homotopic contralateral areas but can differ substantially at small distances along the anterior-posterior axis (Zilles et al., 2015). In line with this, the fact that TEP spectral features diverged more amongst targets that were geometrically closer on the scalp but more different in terms of underlying cortical cytoarchitecture has important implications for the methodological debate on the origin of TEPs. Indeed, this result can be more parsimoniously explained as the effect of a direct cortical activation, rather than in terms of nonspecific sensory responses to scalp or auditory stimulation. These findings support the role for TMS-EEG in the study of electrophysiological correlates of the local structural and functional arrangement of cortical circuits. Importantly, our findings of contralateral TEP symmetry only relate to frequency spectra and not amplitude. TEP amplitude amongst contralateral homologous CAs has been previously shown to be rather asymmetric during the same stimulation session, and this has been considered as a proof of direct cortical activation (Belardinelli et al., 2019). We confirmed this after applying ISP analysis (Casula et al., 2020;Hui et al., 2021)  indices of ISP and IHB by using TMS-EEG, providing the first evidence of correlation between ISP and IHB. In their recent study, they carried out TMS-EEG in 50 healthy participants, stimulating the motor (M1) cortex bilaterally. The resulting TEPs were found to be symmetrical, as measured by ISP and IHB (Casula et al., 2020) whilst these measures showed to be asymmetric in patients with schizophrenia, as demonstrated by Hui et al. (2021). We replicated similar findings for ISP and IHB calculations applied to motor cortices in healthy subjects (Casula et al, 2020), enhancing the reproducibility value of these two indices. Lastly, most studies report stimulation of M1 at or above the RMT to date. Here, we used stimulation intensities below F I G U R E 5 Relative stimulation distances and locations. We used the coordinates from the brain navigation software to calculate the distances between ipsilateral and contralateral homologous target stimulations. (a) Topography of all the targets that we stimulated across participants. The interconnected coloured lines represent the distances we calculated between different targets. Blue represents the distance between left premotor and right premotor targets. Red represents the distance between left motor and right motor targets. The grey colour coding shows the distances between left premotor and left motor, and yellow represents the distance between right premotor and right motor. (b) Box plot of the average of the distances in all the targets we stimulated across all participants. The black line crossing the bars represents the median of each target. The cross inside the bars represents the mean of each target.
the RMT for each motor stimulation, in order to avoid eliciting MEPs, which can contaminate the TEP signal via indirect stimulation of the sensory system (Fecchio et al., 2017).

| Decomposing TEPs with CorrCA
CorrCA identified TEP components that were reliably reproducible across participants and specific to the stimulated region. CorrCA dif-  (Bisogno et al., 2021;Georgopoulos et al., 1982). Further studies are needed to better understand the putative mechanism responsible for this component.

| Monophasic stimulation and hd-EEGtechnical considerations and limitations
We showed that the use of active EEG coupled with a monophasic stimulator can replicate the high-quality TEPs which have been obtained using the traditional methodology reported in the literature, that is, passive electrodes and biphasic stimulator. The use of active electrodes has the advantage of a faster preparation time and is therefore potentially useful in a clinical setting where the duration of the experiment is a critical factor. The feasibility of TEPs recordings with active electrodes in comparison with passive electrodes has already been demonstrated (Mancuso et al., 2021). In this study, active electrodes facilitated fast preparation and reliable maintenance of impedances below 5 kΩ. However, we observed voltage fluctuations after long stimulation sessions, as previously reported (Laszlo et al., 2014).
The use of active electrodes has been associated with more electrical artefact than passive systems, causing a larger decay artefact after the stimulation (Laszlo et al., 2014). We limited the decay artefact by consistently maintaining impedances as low as possible in this study.
Notably, we used online brain navigation instead of standardised coordinates, enabling precise targeting of cortical areas (<2 mm of error). Furthermore, we adopted a real-time TEP visualisation approach , instead of the typical RMT-driven technique, allowing for better control of the necessary intensity, recording of genuine TEPs, and avoidance of muscle and electrical artefacts.
Another potential limitation is represented by the systematic adoption of lower intensities for motor stimulations. Saari et al. (2018) previously showed how TEPs change in relation to the intensity used.
We conducted a correlation analysis between stimulation intensity and natural frequencies across each stimulated area and we did not find any statistical significance (detailed results in Supplementary Material S79.2). This can be ascribed to three main reasons: (1) the difference between the intensities we used was too little; (2) the range of intensities we used (from $90% to $110% RMT) was within a window in which TEP differences are more subtle (in fact the higher differences in TEP shapes in the work by Saari et al. (2018) is found in respect to intensities below 80% of RMT); (3) spectral features are maintained despite changes in the intensities adopted. As such, further work is needed to better understand the impact of TMS intensities on TEP spectral features.  (Fleischmann et al., 2020).

| Future directions
We demonstrate that TEPs are symmetrical in contralateral homologous premotor and motor areas, but asymmetrical in the same areas ipsilaterally, with higher evoked frequencies in the premotor than in motor CAs. Neurophysiological properties measured by TMS-EEG could reflect the functional architecture of the underlying cortical structures and are possibly determined by the underlying neuronal networks. Our findings support the use of TMS as a potential biomarker to assess cortical function in people with lateralized brain pathologies, such as typically seen in many focal epilepsies or stroke.
Our methodology may offer a measure of underlying cortical dysfunction, even in the absence of obvious structural abnormalities, providing an in vivo dynamic assessment of subtle circuit alterations. A normative dataset of TEPs obtained from multiple bilateral cortical targets, with high temporal and spatial resolution, may provide the foundation for future studies investigating focal brain disease and the effect of treatment strategies on a longitudinal basis.


s2fieldsofstudy: Biology Medicine

text:
Highly Efficient Methods to Culture Mouse Cholangiocytes and Small Intestine Organoids

Background Organoids, which enable disease modeling and drug screening closer to an in vivo environment, can be isolated and grown from organs such as the brain, small intestine, kidney, lungs, and liver. To facilitate the establishment of liver and small intestinal organoids, we developed efficient protocols for cholangiocytes and intestine crypts collecting and organoid culturing. Methods Cholangiocytes were collected from intrahepatic bile ducts, the gallbladder, and small intestine crypts by gravity settling and multistep centrifugation methods. The cells isolated were embedded with Matrigel and grew in three-dimensional spheroids in a suitable culture medium. The stability of organoid cells was assessed by subculture, cryopreservation, and thawing. RNA and DNA extraction of organoids, as well as immunostaining procedure, were also optimized. Hand-picking procedures were developed and performed to ensure similar growth characteristics of organoids. Results A large number of cholangiocytes and small intestine crypts were collected under these protocols. Cholangiocytes developed into cyst-like structures after 3–4 days in Matrigel. After 1–2 weeks of cultivation, small intestinal organoids (in-orgs) developed buds and formed a mature structure. Compared to organoids derived from the gallbladder, cholangiocyte organoids (Cho-orgs) from intrahepatic the bile ducts grew more slowly but had a longer culture term, expressed the cholangiocytes markers Krt19 and Krt7, and recapitulated in vivo tissue organization. Conclusions Our protocols simplified the cell collection procedure and avoided the possibility of exposing tissue-derived stem cells to mechanical damage or chemical injury by gravity settling and multistep centrifugation. In addition, our approach allowed similar growth characteristics of organoids from different mammalian tissue sources. The protocol requires 2–4 weeks to establish a stable organoid growth system. Organoids could be stably passaged, cryopreserved, and recovered under protocol guidance. Besides, the organoids of cholangiocytes and small intestines retained their original tissue characteristics, such as tissue-specific marker expression, which prepares them for further experiments such as preclinical in vitro trials and mechanism research studies.

INTRODUCTION
Organoids are in vitro three-dimensional (3D) cultures grown from primarily isolated cells or stem cells (Dutta et al., 2017). Three-dimensional culture enables organoid growth from healthy (Huch and Koo, 2015) or diseased (van de Wetering et al., 2015) human or animal primary tissues. Organoids have potential as preclinical models (Lancaster and Knoblich, 2014;Dutta et al., 2017;Zhang et al., 2021) for drug screening (Kondo and Inoue, 2019) and in mechanistic research (Lancaster and Knoblich, 2014;Vives and Batlle-Morera, 2020). Standardized good manufacturing practice (GMP)-compliant scalable organoids may enable the replacement of damaged human organs (Dossena et al., 2020). What's more, in the widespread spread of world pandemics such as SARS-CoV-2, organoids were also applied to evaluate the intestinal function changes (Zhou et al., 2020), which disclosed a deeper insight into unique gene mutations in gastrointestinal disease (Li, 2021).
As for the cellular composition of organs, the liver mainly comprises hepatocytes, cholangiocytes, and nonepithelial cells (Duncan et al., 2009). The small intestine is almost formed of the intestinal epithelium, which contains intestinal crypt-villus units and Lgr5 + stem cells (Sato et al., 2009), which potentially own stem cell properties. For organoid growth, Matrigel is needed to provide an in vitro 3D environment for stem cell growth. Besides, the proliferation of hepatic and small intestinal stem cells requires epidermal growth factor (EFG), the R-spondin 1 (R-SPO1) protein (notch signaling components) (Sato et al., 2009), growth factors (e.g., N2, B27, and Noggin), hepatocyte growth factor (HGF), and fibroblast growth factor (FGF), as mentioned before.
Based on the above research studies, we developed and improved an efficient method for collection of cholangiocytes and intestinal crypts by gravity settling and multistep centrifugation, which not only simplified the steps of cell separation but also improved the rate of cell acquisition. However, during the experiment, we found a phenomenon that stem cells derived from different mice present different growth characteristics, with some developing into mature organoids in 7-10 days and the others taking more than 10 to develop into mature forms. Therefore, we established an organoid hand-picking procedure (Figure 1) to ensure that the organoids had similar growth characteristics and avoid excessive differences in organoid growth, which is convenient for disease modeling and mechanism research. Under our protocol guidance, the organoids can be passaged, cryopreserved, and thawed, and they are suitable for genetic analysis and immunochemical staining. Compare with other stem cell isolation methods, our procedure does not require additional mechanical equipment (e.g., flow cell sorter) and chemical experiments, like antibody combination, which avoids the possibility of exposing stem cells to mechanical damage Abbreviations: 3D, three dimensional; PSC, primary sclerosing cholangitis; IBD, inflammatory bowel diseases; EGF, epidermal growth factor; FGH, fibroblast growth factor; HGF, hepatocyte growth factor; ROCK, Rhokinase; IF, immunofluorescence; IHC, immunohistochemistry. or chemical injury. Besides, our protocol makes it is easy for beginners to get started and does not require physical exertion. Most importantly, the cell growth rates of stem cells gained from different mammals are unified before further experiments.
Organoids generated from the bile ducts and small intestine have the characteristics of the original tissue, which makes them suitable for disease modeling, drug screening, mechanism research, etc. As recent studies show, organoids may also have a therapeutic investigation potential for diseases such as primary sclerosing cholangitis (PSC) (Loarca et al., 2017), intestinal cancer (Aberle et al., 2018), and inflammatory bowel diseases (IBD) (Meir et al., 2020). Our protocol may provide huge support for the early-stage preparation of such research studies.

Cell Isolation
To isolate bile ducts (cholangiocytes) from mouse liver, the ducts and gallbladder can be digested. In brief, the liver was perfused with PBS 1× through the inferior vena cava before removing the liver tissue. If a small number of cholangiocytes is needed, or if cho-orgs must be cultured rapidly, cholangiocytes can be obtained from mouse gallbladder. After digestion, ducts and crypts from the small intestine can be collected by gravity settling followed by multistep centrifugation. If cholangiocytes need to be counted, bile ducts can be further digested using a single-cell TrypLE Express medium.

Organoid Culture and Establishment of Stable Organoid Lines
To establish a long-term organoid culture, we picked and collected larger organoids to passage and to unify the whole growth speed of organoids in the same culture well. As reported by Huch, when culturing organoids for further experiment, the elements of organoid size, proliferation rate, physical morphology, formation rate, and expression of markers/genes are all needed to be considered (Huch et al., 2013). Therefore, we described an easy hand-picking procedure to achieve this goal. We also provided two methods of cho-orgs culture, one from intrahepatic bile ducts and the other from gallbladder cholangiocytes. These two types of organoids both expressed cholangiocyte specific markers like Krt7 and Krt19, and displayed different transcriptional profiles and differentiation capacity as demonstrated by Casey A. Rimland (Rimland et al., 2021), which provided a basis for selection according to purpose, such as disease simulation or in vitro modeling. Besides, the methods of home-made Wnat3a and R-SPO1 were described by Broutier (Broutier et al., 2016), which were indispensable in organoid culture. When a stable culture line is established, organoids can be maintained for a long time.

Organoid Analysis
To research the gene changes of the organoids, after the removal of Matrigel from the organoid cultures, the procedure of DNA or RNA extraction from the organoids was almost similar with normal cells. Matrigel was removed first using a precold basal medium. It was suggested to use a cell recovery FIGURE 1 | Organoid isolation, culture, growth-rate harmonization, passaging, and freezing. Bile ducts, single bile duct (or gallbladder) cells, and crypts from the small intestine derived from C57BL/6 mice form 3D structures after 7-14 days of culture (5-7 days for gallbladder-derived organoids) in Matrigel. After expansion, the organoids can be characterized or frozen in liquid nitrogen. solution to remove the Matrigel without disrupting organoid structures for immunofluorescence and immunohistochemical staining analysis. Whole-mount immunofluorescence staining enabled 3D structure observation and characterization, such as detecting aimed antibody signal distribution. For observation of cellular morphology and storage of organoids for further staining analysis, paraffin sectioning and immunohistochemical staining are preferred.

MATERIALS Animals
Male or female mice of any genetic background and ranging in age from 6 weeks to 1.5 years (weight 20-28 g) can be used for cho-org culture; for in-org culture, ≤ 6-week-old C57BL/6 mice should be used.

Mouse Liver Digestion Medium
The digestive medium should be prepared fresh and used immediately. Collagenases D and II were dissolved in a sterile washing medium (see below) at a concentration of 0.125 mg/ml and supplemented with 0.1 mg /ml DNase I (dissolved in sterile H 2 O).

Mouse Liver Isolation Medium
The mouse liver isolation medium was a mouse liver expansion medium supplemented with 25 ng/ml recombinant human Noggin, 30% (vol/vol) Wnt3a-conditioned medium, and 10 µM Rho kinase (ROCK) inhibitor (Y-27632 if single cells were cultured). The medium was stored at 4 • C for up to 2 weeks.

Mouse Liver Wash Medium
The DMEM (high glucose, GlutaMAX, and pyruvate) supplemented with 1% FBS and 1% penicillin/streptomycin. The medium was stored at 4 • C for up to 1 month.
F-G clearing solution: 2.5 M fructose and 60% glycerin. Store at 4 • C in the dark for up to 1 month.

I. Culture of mouse cholangiocytes and small intestinal organoids: cell isolation
Mouse liver duct cells may be collected from the liver or gallbladder.
I.i. Collection of bile ducts from mouse liver 1) To collect liver tissue, anesthetize the mouse and expose the liver. Find the inferior vena cava using a sterile swab, place a sterile cotton ball beside the liver, inject PBS 1× using a 10-mL injector, and cut off the portal vein immediately when the liver swells. By standard surgical procedures, remove the liver as an entire organ and transfer it to a 10-cm Petri dish. 2) Place the Petri dish on ice and transfer it to a biological safety cabinet. Preheat the digestion solution to 37 • C. 3) Cut the liver tissue into small pieces (<1 mm 3 ) using fine scissors. Transfer the small pieces to a 50-mL sterile centrifuge tube, add up to 5 ml precooled washing medium, and pipette it up and down several times using a 10-ml pipette to wash the minced tissue. Repeat the washing procedure. 4) Transfer the tissue to a new 50-ml sterile centrifuge tube.
Add 5 ml pre-warmed digestion medium. Shake the tube at 120-160 rpm and 37 • C for ∼2 h. 5) During incubation, check the appearance of bile ducts using a light microscope. Pipette up and down the supernatant in a biological safety cabinet using a 1-ml pipette, and transfer ∼200 µl of the solution to a glass slide for observation ( Figure 2B, upper row). If no bile ducts are present, return the solution to the shaker. Perform the checking at 20-30 min intervals. Ducts usually appear after 60 min. 6) When bile ducts appear, transfer the digestion supernatant to a fresh 50-ml centrifuge tube, add the same volume of precooled washing medium, and centrifuge at 80 g for 4 min at 4 • C. Discard the supernatant and add 15 mL precooled washing medium; repeat the centrifugation to remove the remaining digestion solution. 7) Add the precooled washing medium to the pellet and pipette it up and down to mix. Place the 50-mL centrifuge tube upright on ice for 30 min. 8) Remove the supernatant without disrupting the pellet. Add 5 ml precooled washing medium, transfer the mixture to a fresh 15-ml centrifuge tube, and centrifuge at 60 g for 2 min at 4 • C. 9) Remove the supernatant carefully. Add 1 ml precooled basal medium, transfer the mixture to a 1.5-ml microcentrifuge tube, and centrifuge at 500 rpm for 2 min at 4 • C. 10) The pellet can be directly cultured. If there is a need to collect single bile cells, follow the next procedure.
I.ii. Enrichment of single cholangiocytes 1) Resuspend bile ducts in 5 ml prewarmed TrypLE solution supplemented with 5 µl DNase I (10 mg/ml). Using narrow 1,000 µl tips, pipette the solution up and down to mix and incubate at 37 • C for 2-10 min. 2) Check the solution every 2 min using a bright-field microscope. Stop the digestion when majority (85-95%) of the mixture consists of single cells by adding the precooled wash medium (Figure 2A, upper row). 3) Add 10 ml cold washing medium to stop the digestion.
Transfer the mixture to a 50-ml centrifuge tube through a 70-µm filter and centrifuge at 300-350 g for 5 min at 4 • C. Remove the supernatant and repeat the centrifugation to wash out any remaining TrypLE. Add 1-5 ml washing medium to resuspend the pellet. 4) Enumerate the cells using a standard cellcounting chamber.
I.iii. Collection of cholangiocytes from gallbladder Prewarm the mouse liver digestion medium and TrypLE solution (supplemented with 0.1% [vol/vol) DNase I [10 mg/ml]) to 37 • C. 1) By standard surgical procedures, anesthetize the mouse and expose the liver. Strip the gallbladder using two ophthalmic forceps and transfer it to a 6 mm dish containing precooled mouse liver washing medium. 2) Cut the gallbladder into small pieces using ophthalmic scissors, transfer the pieces to a 15-ml sterile centrifuge tube using a 1,000 µl Eppendorf pipette, add ∼5 ml precooled washing medium, gently pipette the solution up and down, and centrifuge it at 4 • C, 200-250 g, for 4 min to remove bile. 3) Discard the supernatant and add 5 ml mouse liver digestion medium. Shake the tube at 120-180 rpm and 37 • C for 1 h. 4) Add 5 ml precooled washing medium to stop the digestion, centrifuge at 4 • C and 200-250 g for 4 min and discard the supernatant. 5) Resuspend the gallbladder tissue in 1 ml prewarmed TrypLE solution and pipette it up and down 30 times to isolate single cholangiocytes. 6) Incubate the tube for 2-4 min in a 37 • C culture bath and pipette it up and down 30 times. 7) Add t∼5 ml precooled wash medium and filter the mixture through a 70 µm mesh into a 50-ml centrifuge tube using a 1,000 µl Eppendorf pipette. Pipette the fluid into a 15ml centrifuge tube, and centrifuge at 4 • C and 300-350 g for 4 min. 8) Discard the supernatant and enumerate the cells.
The plates are ready for Matrigel embedding and 3D organoid culture.
I.iv. Collection of crypts from mouse small intestine 1) Anesthetize the mouse; by standard surgical procedures, expose and remove the intestine using ophthalmic forceps. Cut the optional intestine segment 1 cm above the end of the ileum and 2-3 cm below the stomach, and transferred them to a 10 cm dishes containing PBS 1×. 2) Put the 10-cm dish on ice. Squeeze out the intestinal contents using forceps and cut the tissue longitudinally. Use a blunt instrument (e.g., the curved part of curved dissecting forceps) to scrape the intestinal villi, wash twice or thrice and cut the tissue into 1-2 cm pieces.
3) Transfer the pieces to a 50 ml centrifuge tube and add 20 ml mouse intestine digestion medium. Put the tube on a shaker at 120-160 rpm and 4 • C and incubate for about 30 min. 4) Carefully remove the supernatant and resuspend the tissue in a new 50 ml centrifuge tube, add ∼25 mL PBS 1×, and vigorously pipette the solution up and down 30-50 times to isolate crypts from the tissue. 5) Filter the supernatant through 100-and 70-µm meshes into a new 50 ml centrifuge tube. 6) Allow the tube to stand for 8 min and discard the supernatant or transfer them to a new tube for another 8 min-standing circulation. Gather the plates together and check the proportion of the crypts. Add 1-5 ml PBS 1× to the white sediment and pipette it up and down gently.
Observe the mixture under a light microscope. 7) If the proportion of single cells is too high, add ∼25ml precooled PBS 1×, and repeat the 8-min standing circulation procedure in step (D-6) to collect a higher proportion of crypts. 8) Count the crypts using a counting board. Incubate for 15-20 min at 37 • C until Matrigel solidifies. 4) Add the appropriate medium to each well (500 µl per well for a 24-well plate or 250 µl per well for a 48-well plate) (liver isolation medium for cho-org culture, intestine expansion medium for in-org culture). 5) Incubate the plates at 37 • C in a 5% CO 2 atmosphere. For cho-orgs, after 3 days, change the isolation medium with the expansion medium and incubate for ∼14 days. For in-orgs, retain the intestine expansion medium. Change the medium every 3-4 days. Organoids will start to develop on days 3-5.
III. Organoid passage 1) Prewarm culture plates for 1 h-overnight. Place Matrigel on ice and thaw before use. Prewarm the TrypLE Express solution (∼2 ml per tube) in a water bath at 37 • C. 2) To disrupt the basal matrix, add 500-1,000 µl precooled basal medium (for cho-orgs) or PBS 1× (in-orgs), and pipette the mixture up and down using a 1,000 µL pipette. Transfer the organoid suspension (three wells for 24-well culture plates or six wells for 48-well culture plates) to a 15 ml centrifuge tube, add ∼10 ml precooled basal medium (cho-orgs) or PBS 1× (in-orgs) to the top, and mix by pipetting vigorously 5-10 times to wash away Matrigel.
3) Centrifuge the tube at 200-250 g for 5 min at 4 • C. 4) Discard the supernatant, leaving 100 µl mixture. 5) Add the prewarmed TrypLE Express solution and mix by vigorously pipetting up and down using a 1,000 µl pipette. Transfer the tube to a water bath at 37 • C for 1-4 min, checking every 2 min under a light microscope. When majority (85-95%) of the material is single cells, add ∼10 ml precooled basal medium to stop digestion. 6) Centrifuge the tube at 300-350 g for 4 min at 4 • C, and carefully aspirate the supernatant. 7) Organoids can be mechanically dissociated and split at a 1:3-1:6 ratio. Resuspend the cells in Matrigel (40-50 µl per well for 24-well plates or 20-25 µl per well for 48well plates). Pipette the mixture up and down gently to resuspend the cells. Add a droplet of the mixture to the center of each well. Incubate for 15-20 min to allow for Matrigel polymerization. 8) Overlay the cultures with the expansion medium (500 µl per well for 24-well plates or 250 µl per well for 48well plates). 9) Replace the medium every 2-3 days.

IV. Harmonizing the growth rates of organoids
Organoids developed from different primary cells grew at different rates, for some experimental needs, organoids in different growing state are hard to perform experiments. To resolve this problem, we purified cho-orgs with similar growth rates using the procedure below, which can be repeated once to thrice according to growth situation. In addition, intestinal organoids can also use the following method to uniform growth rate.
1) When cho-orgs have budded for 2-3 days (5-7 days after seeding), check their growth state daily under a light microscope. 2) Before cells aggregate in the larger cho-orgs, prepare to selectively passage then, as mentioned in III-1, prewarm the culture plates and TrypLE Express solution, and thaw the Matrigel before use. 3) Move the plates to a sterile environment. Under a light microscope, transfer the larger cho-orgs to the precooled basal medium using a 10-µl pipette. 4) Subsequent steps are the same as for organoids passage; please follow steps III-3-8.
V. Cryopreservation and thawing of organoids V.i. Freezing organoids 1) At least one confluent well (24-well plate) or two confluent wells (48-well plates) of organoids are needed per cryovial tube. 2) Proceed as in steps (III-1-5) to digest the organoids into single cells and resuspend the cells in 500 µl precooled freezing medium per well (24-well plates) or two wells (48well plates). Transfer the mixture to cryovials (500 µl each), and immediately place them on ice. Transfer the cryovials at −80 • C and then to liquid nitrogen after 24 h. The organoids can be stored for years.
V.ii. Thawing of organoids 1) Prewarm a 15 ml tube with 10 ml basal medium (for choorgs or in-orgs) at 37 • C. Prewarm a 24-or 48-well plate according to need. 2) Incubate the cryovial in a 37 • C water bath and remove when the frozen cell mass is almost completely thawed. Transfer the thawed cell aggregates to the prewarmed basal medium and pipette it up and down gently. 3) Centrifuge the tube at 250-300 g for 4 min at 4 • C. 4) Remove the supernatant without disturbing the pellet. 5) Proceed as in steps (II-1-3) to seed the cells in Matrigel and add the appropriate expansion medium (500 µl per well for 24-well plates or 250 µl per well for 48-well plates) to each well. 6) Replace the medium every 2-3 days.

VI. Analysis of organoids
To characterize the organoids, use option (VI.i) for immunofluorescence analysis, (VI.ii) for immunohistochemical staining, and (VI.iii-iv) for isolation of DNA (iii) or RNA (iv), VI.i. Immunofluorescence staining 1) To collect the organoids completely, remove the expansion medium, and add 500-1,000 µl precooled Cell Recovery Solution to each well. 2) Gently shake the plate horizontally at 4 • C for 30-60 min to disrupt Matrigel. 3) Cut off the top of 1,000 µl tips. Blow the tips twice in precooled 1% (v/v) PBS-BSA, and wash 15 ml centrifuge tubes using 1% (v/v) PBS-BSA to precoat the tips and tubes. 4) Transfer the mixture to the precoated tubes using the precoated tips, add PBS 1× to ∼10 ml and mix gently. Centrifuge at 70 g and 4 • C for 4 min, and carefully remove the supernatant. 5) Repeat the PBS 1× wash steps once or twice to wash out Matrigel completely. 6) Fix the organoids by resuspending in 4% (w/v) paraformaldehyde at 4 • C for 45 min and mix once or twice. 7) Add ∼10 ml precooled 0.1% (vol /vol) PBT, mix, and centrifuge at 70 g and 4 • C for 4 min. 8) Remove the supernatant and resuspend the organoids in the precooled washing buffer. Transfer the mixture to a 24well plate (>200 µl per well) and incubate at 4 • C for 15 min. 9) When the organoids sink to the bottom, tilt the culture plate at 45 • and remove the washing buffer to leave about a 200-µl liquid. 10) Add a two-fold concentration of primary antibody (diluted in 0.5% PBS-BSA) (200 µL) to each well. Incubate overnight at 60 rpm and 4 • C. 11) Add 1 ml washing buffer to each well and pipette gently. 12) When the organoids sink to the bottom (3 min), remove 1 ml washing buffer, add 1 mL washing buffer, and incubate for 30 min. 13) Repeat step (11) at least twice. 14) When the organoids sink to the bottom, remove the washing buffer to leave a 200 µl liquid. 15) Add a two-fold concentration of secondary antibody (diluted in 0.5% PBS-BSA) (200 µl) to each well. Incubate overnight at 60 rpm and 4 • C. 16) Repeat 10-13. Transfer the organoids to 1.5 ml EP tubes and centrifuge at 70 g and 4 • C for 4 min. The organoids can be stored in the washing buffer at 4 • C for 2 days. 17) Remove the washing buffer without touching the plates. 18) Add the F-G clearing solution (>50 µl, RT) to the EP tube using top-cut 200-µl tips. 19) Add an appropriate volume of DAPI and incubate for 20 min at room temperature. The organoids can be stored in the F-G clearing solution for 1 week at 4 • C or 6 months at −20 • C. 20) Transfer the organoids to a 24-well plate using 200-µl top-cut tips. 21) The organoids can be subjected to immunofluorescence imaging.
VI.ii. Immunohistochemical staining of organoids 1) To obtain the organoids completely, perform steps VI-C 1-6. 2) Remove 4% (w/v) paraformaldehyde, add ∼10 ml precooled PBS 1×, and centrifuge at 70 g and 4 • C for 4 min. 3) Repeat the washing step once or twice. 4) Remove the supernatant and add ∼10 ml 70% alcohol. The organoids can be stored at 4 • C in 70% alcohol for 1 week. 5) Adjust the water bath to 65 • C and prewarm Histowax (∼3 ml per sample), plastic straws, soft EP tubes, and a metal mold. Be careful to keep the materials dry. 6) Place a centrifuge tube on ice upright until the organoids sink to the bottom. Remove the supernatant, resuspend the organoids in 0.5% (w/v) eosin solution to dehydrate, and stain for >30 min. 7) When the organoids sink to the bottom, discard the supernatant and resuspend in 100% alcohol to wash for at least 30 min. Repeat these wash steps thrice. 8) Resuspend the organoids in dimethylbenzene and wash thrice for ∼30 min each. 9) Remove the dimethylbenzene. 10) Place the centrifuge tube containing the organoids mentioned above in a 65 • C water bath, absorb appropriate volume of prewarmed liquid Histowax (≤500 µL) to the organoids using the prewarmed plastic straws, quickly pipette twice or thrice, and transfer the mixture to the prewarmed soft EP tubes. 11) Incubate the soft EP tubes at 65 • C overnight. 12) Transfer the soft EP tubes to RT or a cold area to solidify Histowax. 13) Peel off the tube carefully and cut the Histowax to a smaller size suitable for embedding. This step concentrates the organoids, facilitating their staining and visualization.
14) Re-embed the small wax block to a new Histowaxin metal mold. 15) Cool the mold and take out Histowax, which can be stored at RT for years. 16) Cut Histowax into pieces for further staining.
VI.iii. Isolation of DNA 1) Perform the procedure in main text III 2-3 to remove Matrigel. 2) Discard the supernatant, add ∼1 ml precooled PBS 1×, and pipette the mixture up and down thrice. Transfer the mixture to 1.5 ml EP, and centrifuge at 250 g and 4 • C for 4 min. 3) Resuspend the organoids in PBS 1×, transfer them to a 1.5 ml EP tube, and centrifuge at 300 g and 4 • C for 4 min. 4) Discard the supernatant and add 30 µl Direct-PCR solution and 0.5 µl proteinase K per sample. 5) Incubate at 60 • C overnight. 6) Centrifuge at ≥8,000 g at RT for 10 min. 7) DNA can be stored at 4 • C. 8) If necessary, purify the DNA using a DNA purification kit.
VI.iv. Isolation of RNA 1) To collect the organoids, follow steps VI.iii 1-3 to collect cell mass. 2) Remove the supernatant, add 350 µl RLT to each 1.5 ml microcentrifuge tube, and vortex-mix for 30 s. 3) To promote cell mass lysis, draw the mixtures at least 5 times without RNase using a sterile syringe fitted with a no. 20 needle. 4) The mixtures can be stored at −80 • C for 1 month. 5) RNA can be extracted following the RNeasy Mini Kit (Qiagen, Hilden, Germany) protocol guides.

Troubleshooting
Troubleshooting advice can be found in Table 1.
Step Times Step I. i-ii: collection of mouse liver bile-duct cells: 4 h.
Step I. iii: collection of mouse gallbladder cholangiocytes: 2 h.
Step I. iv: isolation of mouse intestinal crypts: 2 h.
Step II: seeding of mouse cholangiocytes or crypts: 30 min.
Step V: cryopreservation and thawing of mouse organoids: 30 min each.
Step VI: analysis of organoids: 2 days for isolation of DNA, 40-60 min for isolation of RNA, 3 days for immunofluorescence staining, and 2 days for immunohistochemical staining.

Establishment of Organoid Cultures
The bile ducts, cholangiocytes, and intestine crypts embedded in Matrigel developed into 3D structures (Figure 2). The choorgs from cholangiocytes were single cystic spheres, while the primary cho-orgs from bile ducts comprised multi-cystic spherelike organoids but can be separated after passaging and grow into FIGURE 3 | Development of cho-orgs and in-orgs. Cholangiocytes (bile ducts) and crypts from the small intestine embedded in Matrigel developed into cyst-like structures. (A,B) Cells congregated in the cysts of in-orgs (day 3) and were outside the cho-orgs (day 4); after culturing for 10-13 days, the cho-orgs (day 10) and in-orgs (day 13) increased in size, and the in-orgs developed three buds (white arrow) to form a mature structure (bar, 100 µm). (C) Mature in-orgs after culturing for 7 days after passage, they were bud-shaped and had aggregated cell clumps in the organoids (bar, 100 µm).
single cystic spheres, as in the case of cho-orgs derived from single cholangiocytes. Besides, the In-orgs were bud-shaped and had aggregated cell clumps in the organoids.

Growth of Organoids
After seeding for 2-4 days, primary stem cells from the cholangiocytes or small intestine started to develop into cyst-like structures ( Figure 3A). After 10-14 days, the organoids increased in size and matured ( Figure 3B) and could be passaged at a 1:3-1:4 ratio for further expansion (Figures 2A, 3C).

Characteristics of Cho-Orgs and In-orgs
Cho-orgs expressed the cholangiocytes markers Krt19 ( Figure 4A) and Krt7 (Figure 4B), showed high proliferative activity, and consisted of more than one cell layer ( Figure 4C). Immunofluorescence staining for CK7 (red) in a 3D structure (bar, 100 µm). (C) The cho-orgs consisted of more than one cell layer and express the proliferation marker Ki67 (bar, 100 µm). (D) The cho-orgs from intrahepatic bile ducts were cultured for 7 days after seeding and passaged seven times for a total culture duration of >64 days (bar, 200 µm).
We found a phenomenon that the cho-orgs from the intrahepatic bile ducts grew slower than the cho-orgs from gallbladders. Formation of mature organoids, about 200 nm in size, took only about 5 days for gallbladder-derived bile duct cells compared to the 7-14 days for primary intrahepatic bile duct cells. However, the latter could be passaged for a longer period ( Figure 4D).
Besides, the in-orgs expressed the enterocyte cell marker CK20 before they matured (Figure 5A), and the cell connections were stable as shown in the complete tight junction protein CLAUDIN-1 staining results ( Figure 5B). In addition, the intestinal permeability of the in-orgs was proved to be normal, and the fluorescent dye did not enter the cavity of the organoid, as identified by FITC-dextran-4KD (FD4) staining analysis ( Figure 5C).
It is worth mentioning that the digestion time of the organoids in TrypLE Express solution is crucial for the viability of the next generation of organoids during passaging. On  the one hand, prolonged incubation time leads to overdigestion and lower cell viability; on the other hand, insufficient digestion can lead to decrease in the number of organoids. Therefore, the digestion situation of organoids should be monitored under a light microscope. The optimum time of digestion can be determined when 80-90% of cells are single cells.

Stable Growth of Cho-Orgs After Cryopreservation and Recovery
The organoids grew stably after thawing, as observed by light microscopy (Figure 6). After stable culture, homogenization of growth rate manipulation, the organoids were ready for RNA/DNA isolation, immunochemical staining, cryopreservation, or further experiments.

CONCLUSIONS
Our protocols have high repeatability if tissue dissociation steps are performed correctly, and the composition of medium is precise. Besides, we simplify the cell collection procedure by gravity settling and multistep centrifugation to collect cholangiocytes/bile duct and intestine crypts from mouse liver cholangiocytes or small intestine, although some unwanted cells may also be collected during the process, which will disappear gradually after 1-2 passages of the organoids. In addition, when experiments are operated properly, the possibility of exposing stem cells to mechanical damage or chemical injury will be avoided. After passaging, cryopreservation, and thawing, the organoids are in a stable growth state and suitable for mechanistic analysis, preclinical research, drug screening or further experiments.

DATA AVAILABILITY STATEMENT
The original contributions presented in the study are included in the article/Supplementary Material, further inquiries can be directed to the corresponding author/s.

ETHICS STATEMENT
The animal study was reviewed and approved by the First Affiliated Hospital, College of Medicine, Zhejiang University (reference number 2020-1088).

AUTHOR CONTRIBUTIONS
WC and HC contributed to the design of the study, method optimization, and writing of the manuscript. WC, QY, and RW contributed to organoid culture and experiments. WC, BF, JC, YX, and JY contributed to method optimization. WC and QY contributed to the production of Figures. LL conducted study supervision. All authors have read and approved the final version of the manuscript.