# Covalent anchoring of tethered ligands to chemogenetic handles for targeted in vivo neuropharmacology

Maria Ciscato, Kian Stenbolt, Alexandre Mourot, Nicolas Guyon

► To cite this version:

Maria Ciscato, Kian Stenbolt, Alexandre Mourot, Nicolas Guyon. Covalent anchoring of tethered ligands to chemogenetic handles for targeted in vivo neuropharmacology. STAR Protocols, 2025, 6 (4), pp.104186. 10.1016/j.xpro.2025.104186. hal-05516845

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https://hal.science/hal-05516845v1

Submitted on 18 Feb 2026

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# Covalent anchoring of tethered-ligands to chemogenetic handles for targeted *in vivo* neuropharmacology

Maria Ciscato,<fnref n="1" /> Kian Stenbolt,<fnref n="1" /> Alexandre Mourot,<fnref n="1" />,<fnref n="2" />,<fnref n="3" />,* and Nicolas Guyon<fnref n="1" />,<fnref n="2" />,<fnref n="4" />,**

## Summary

Covalent anchoring of tethered-ligands to chemogenetic handles (CATCH) is a chemogenetic approach that enables sustained, localized and pharmacologically-specific receptor antagonism. Here, we present a protocol for designing and applying CATCH in mice. We describe how to integrate CATCH with *ex vivo* and *in vivo* electrophysiological recordings, as well as detail procedures for combining CATCH with behavioral assays. CATCH enables the specific control of neurotransmitter receptor function, offering a tool for both neurophysiology and behavioral studies.

For complete details on the use and execution of this protocol, please refer to Jehl, Ciscato, Vicq, Guyon et al.<fnref n="1" />

## Graphical abstract

[To be inserted here]

## Before you begin

Precise manipulation of endogenous receptors *in vivo* remains a significant challenge in neuroscience, particularly in the context of physiology and behavior, due to the lack of specific pharmacological tools and the difficulty in achieving sustained and localized effects. The application of traditional pharmacological tools is particularly problematic in *in vivo* electrophysiology, as compounds must be pressure-injected during experiments, often prematurely ending the recording.

<footnote n="1">¹Brain Plasticity Unit, ESPCI Paris, PSL University, CNRS, 75005 Paris, France</footnote>

<footnote n="2">²Senior author</footnote>

<footnote n="3">³Technical contact</footnote>

<footnote n="4">⁴Lead contact</footnote>

<footnote>*Correspondence: alexandre.mourot@inserm.fr</footnote>

<footnote>**Correspondence: nicolas.guyon@espci.fr</footnote>
<figure><img src="image_2.png" /></figure>

The CATCH approach overcomes these limitations by enabling sustained, localized, and receptor-specific antagonism through a chemogenetic strategy, allowing for precise and long-lasting manipulation of neuronal signaling. This protocol is compatible with a variety of downstream applications, including *ex vivo* and *in vivo* electrophysiology and behavioral assays such as conditioned place preference, and can be adapted to other receptor systems with appropriate genetic targeting. For optimal reproducibility and interpretation, careful attention to experimental controls and validation steps is essential.

Innovation

CATCH is a chemogenetic approach that enables sustained and receptor-specific antagonism *ex vivo* and *in vivo*. CATCH combines genetic receptor modification with a tethered ligand for irreversible binding. The genetic modification consists in introducing a cysteine residue that selectively reacts with maleimide-tethered ligands. CATCH is an extension of the photoswitchable tethered ligand (PTL) approach, in which light-sensitive ligand are covalently attached to cysteine-substituted ion channels or receptors.²⁻¹⁶ While PTLS offer reversible and rapid control of protein function, their *in vivo* use usually requires local light delivery within target brain regions. This can be technically demanding and often necessitates tethering animals to an optical fiber, which restricts movement. Moreover, many experimental settings do not require rapid on/off switching, and instead benefit from prolonged receptor antagonism. We demonstrated that CATCH achieves long-lasting (several days) and efficient (>80%) receptor blockade,¹ making it particularly suitable for neurophysiological and behavioral studies. A key advantage of CATCH is the minimal genetic modification, which preserves native receptor pharmacology and allows investigation of receptor function within intact neural circuits. Although the cysteine substitution is not strictly bioorthogonal, no measurable off-target effects have been observed.¹ This is provided that proper wild-type controls with the tethered ligand are included. This strategy has also been successfully applied to other systems, such as potassium channels, using cysteine-reactive quaternary ammonium groups¹⁷ and peptide toxins¹⁸, further illustrating the versatility of this approach. Likewise, covalent activators could be deployed instead of antagonists, although this approach is limited to non-desensitizing receptors or ion channels.¹⁹

Institutional permissions

All experiments on mice must first be approved by the appropriate institutional authority in accordance with the national guidelines. Our protocol was approved by the French Ministry of Education in Research and Paris Sciences et Lettres (PSL) University after consultation with the ethical committee #59 in accordance with the European Directive 2010/63/EU of the European Parliament.

Design of the receptor-ligand pair

CATCH is a chemogenetic method that relies on a uniquely selective receptor-ligand pair, in which a reactive antagonist is covalently tethered to a genetically-modified receptor bearing a strategically placed cysteine substitution. This engineered cysteine residue is positioned on receptor surface, in close proximity to the agonist binding site, to enable site-specific, irreversible antagonism upon covalent reaction with the ligand (Figure 1A).

1. Design of the ligand. The tethered antagonist is made up of three elements: a cysteine-reactive, maleimide head group; a flexible, water-soluble polyethylene glycol (PEG) linker; and a functional ligand (antagonist). Specifically, for manipulating nAChRs, we designed MPEG₄Ch
84 (maleimide - tetraethylene glycol - choline), which carries a four glycol unit PEG linker and a
85 choline headgroup as an antagonist (Figure 1B).

86 a. Reactive head group: maleimides remain the preferred cysteine-reactive group due to their excellent reactivity (they react within minutes with high yields),²⁰ their biocompatibility and their commercial availability. Furthermore, maleimides-PEG linkers show excellent stability after bioconjugation,²¹ a key feature for irreversible antagonism.

91 b. Linker: The length of the PEG linker is critical: a short linker (e.g., four units as in MPEG₄Ch) ensures that the antagonist remains concentrated in the immediate vicinity of the binding pocket, thereby increasing local efficacy while minimizing off-target interactions. We recommend keeping the linker as short as possible.

95 c. Pharmacophore: There is a tradeoff between using a minimal pharmacophore (the smallest chemical structure required for biological activity) and a high-affinity ligand. High affinity ligands improve binding specificity and may reduce side effects, but can be synthetically demanding. In contrast, minimal pharmacophores (e.g., the triethylammonium group in MPEG₄Ch) benefit from facile synthesis, and their weak intrinsic binding is offset by covalent tethering, which ensures high local concentration.

102
103 **Note:** Custom synthesis services are available from several companies. For example, Broadpharm® offers a wide range of maleimide-PEG derivatives with various headgroups (Figure 1C) —including alcohols, amines, esters, amino acids, and alkynes (the latter enabling click chemistry for further modification). It is important to use compounds of high purity; In our case, Broadpharm® specifies a purity of 98% for MPEG₄Ch Chloride powder.

108 2. Design of the cysteine-substituted receptor: On the receptor side, the cysteine mutation should be introduced at a surface-exposed position near the agonist binding site, but without altering receptor function in its unbound state. Identifying an optimal labeling site requires detailed knowledge of receptor structure and of the structure–activity relationships (SAR) of known ligands. For nAChRs, high-resolution structural models were previously used to design attachment points for photoswitchable ligands.⁵,¹¹ The design of MPEG₄Ch was guided by the structure of a previously developed photoswitchable antagonist, ensuring compatibility with existing cysteine-engineered nAChRs and expanding toolset versatility.¹ We selected glutamate 61 (E61) in the β2 and β4 nAChR subunits, located at the extracellular entry of the acetylcholine binding pocket (Figure 1D). This mutation preserves native receptor function prior to ligand conjugation.¹,⁵ Below we provide a general strategy for cysteine site selection.

119 a. Identify candidate residues: Download a high-resolution structure from the Protein Data Bank (https://rcsb.org), ideally with a docked ligand. For nAChRs, the human α3β4 nAChR with nicotine (PDB 6pv7)²² is suitable (Figure 1D). Because tethered ligands are flexible, complex docking is not required; instead, compute the distance between the docked ligand and candidate side chains using visualization software (e.g., VMD, https://www.ks.uiuc.edu/Research/vmd/).

125 b. Filter candidates: Choose residues that are not essential for receptor function. Multi-sequence alignment (Clustal Omega https://www.ebi.ac.uk/jdispatcher/msa/clustalo?stype=protein) can help exclude highly conserved residues (marked * or :) that are more likely to be functionally critical.
129 Protein sequences in the FASTA format can be retrieved from the NIH database (https://www.ncbi.nlm.nih.gov/protein/).

130 c. Introduce cysteine mutation: This can be performed with standard mutagenesis kits or outsourced to plasmid synthesis companies.

131 d. Validate receptor function. Mutant receptors should be tested in functional assays (e.g., electrophysiology for nAChRs) to confirm that activity is preserved.

132 e. Verify irreversible blockade. Once the covalent reaction is complete and excess reagent is washed out, the mutated receptors should be irreversibly blocked, with no further antagonist required to maintain the blockade, while WT receptors should remain unaffected (Figure 1E).

139 **Note:** Comparison with other chemogenetics methods.

140 A direct variant of the CATCH method involves photoswitchable tethered ligands (PTLs), whose geometry can be modulated by light, enabling rapid and reversible control of protein function.²⁻¹⁶ These photoswitchable systems are particularly powerful for modulating synaptic transmission with high temporal precision. However, their application in behaving animals remains technically challenging due to the difficulty of delivering both the ligand and light locally. Moreover, they are not well suited for long-term experiments, as they often require mice to be tethered to external light sources, limiting behavioral flexibility. However, it should be noted that in the absence of illumination, some PTLs, the ones that bind to receptors in their dark-adapted state, act much like CATCH, producing sustained antagonism through covalent binding.²,⁴,⁹,¹²,¹⁵ However, this is not the case for light-controllable nAChRs, in which light is required to present the ligand in its binding pocket.⁵,¹¹,¹³ Thus, while light delivery can present practical challenges in behaving animals, this does not diminish the fundamental advantage of PTLs in offering temporally precise, reversible control when optical stimulation is used.

152 The use of genetically introduced cysteine residues presents several advantages: it enables efficient and selective labeling with maleimides and generally results in minimal perturbation of receptor function. However, this strategy is not fully bioorthogonal, as endogenous cysteine residues are present on native proteins. As a result, non-specific binding of the ligand may occur. Although no off-target effects were observed in our study¹ or with PTLs,⁶,¹³,¹⁴ it remains essential to perform control experiments using the tethered ligand in non-transfected or native tissue. To enhance labeling specificity, alternative tagging strategies have been developed.²³ In particular, SNAP and CLIP tags offer high specificity for labeling genetically modified proteins.²⁴⁻²⁶ However, these tags are relatively large protein domains (~20 kDa), and their fusion to membrane receptors may interfere with receptor folding or function. In the specific case of nAChRs and similar cys-loop receptors (i.e. GABA<sub>A</sub>, glycine and 5HT<sub>3</sub> receptors), incorporation of tags at the N- or C-terminus alters receptor expression and activity,²⁷ rendering this tagging approach inapplicable for this receptor class.

164 An entirely different chemogenetic strategy—the “bump-hole” approach—involves engineering a mutation (hole) in the receptor's ligand-binding pocket, paired with a modified synthetic ligand (bump) that only fits the altered site.²⁸ This approach has been successfully used to develop engineered GPCRs, including the widely known DREADDs (Designer Receptors Exclusively Activated by Designer Drugs),²⁹,³⁰ as well as cys-loop receptors³¹ that are activated selectively by synthetic ligands. While effective for remote and specific control of neuronal activity, these systems rely on receptors that no longer respond to their endogenous ligands, and are therefore unsuitable for studying the native physiological roles of these receptors in intact circuits.²³
172 Design of knock-in mice or viral vectors

173 Two knock-in (KI) mouse lines, β2E61C and β4E61C, were generated by targeted homologous recombination to introduce a single point mutation into the endogenous Chrnb2 or Chrnb4 gene. In these mice, all native β2 or β4 subunits are replaced by cysteine-substituted versions, ensuring unaltered receptor density, localization, and regulation under native promoters. This strategy preserves physiological expression patterns, but has a major limitation: the cysteine substitution is present in all cells expressing the targeted subunit, making it impossible to restrict covalent ligand labeling to specific cell types. Anatomical specificity can instead be achieved by local ligand infusion, though this does not provide cell-type resolution.

181 To achieve anatomical specificity, ligand delivery is performed locally, via stereotaxic infusion. However, a more refined approach would involve cell-type-specific replacement of endogenous nAChR subunits with cysteine-substituted variants. Although such a method is not yet widely established, recent advances in gene editing techniques offer a potential path forward.³²

185 **Note:** The KI mouse lines were generated on a C57BL/6N background by homologous recombination at the Phenomin core facility (Institut Clinique de la Souris, Illkirch, France). Exons 3 and 4 were fused to prevent alternative splicing at the E61 codon/splice acceptor site. Knock-in mice were bred as homozygotes.

189 **Alternatives:** Knock-in mice can be produced by various academic or commercial facilities, such as GenOway (France), Polygene (Switzerland), Charles River (Global), or Cyagen (Global).

191 An alternative strategy is to deliver cysteine-mutated receptors via viral vectors. For instance, we used a lentivirus encoding pGK-β2E61C-IRES-eGFP to express β2E61C receptors (available at Addgene, see Key resource table for more information) to target expression of β2E61C receptors in the ventral tegmental area.¹³ The single cysteine mutation E61C was inserted into pLenti-pGK-β2-IRES-eGFP by site-directed mutagenesis using the Quickchange II XL kit (Agilent). Mutations were verified by DNA sequencing.

196 This viral approach can be combined with MPEG4Ch, and offers crucial advantages such as restricting mutant expression to specific structures or cell types, thereby providing unparalleled manipulation of pre- versus postsynaptic receptors. We did not detect any change in overall nicotine-induced currents in transduced cells, suggesting no alteration in receptor number.¹³ This outcome likely reflects the fact that β2 subunits form obligate heteromers with α subunits, helping maintain total receptor numbers in balance. However, viral expression does not guarantee complete replacement of endogenous subunits, and stoichiometric alterations (two vs three β subunits per pentamer) could still affect receptor sensitivity to acetylcholine or nicotine. More generally, caution is warranted when applying this approach to homomeric receptors (e.g., α7 nAChRs), since ectopic expression may change total receptor number and complicate interpretation. In such cases, promoter strength becomes an important parameter to adjust expression levels and maintain physiological receptor density. Finally, viral strategies add technical requirements, including the need for an additional transduction step and thorough validation of receptor numbers.

210
211 **Note:** When selecting and ordering viral vectors, always include a control virus. The control virus should express only the fluorophore and match the experimental virus in serotype, promoter, and recombinase dependence.

214 215 **Note:** Upon receipt, thaw the virus and prepare aliquots. Store aliquots at -80°C for long-term stability, and use only one aliquot per experiment to avoid repeated freeze-thaw cycles. Thaw aliquots on ice before use and do not refreeze. AAV aliquots can be kept at 4°C for up to 6 months; lentivirus should be used immediately.

219 Key resources table

220

<table><thead><tr><th>REAGENT or RESOURCE</th><th>SOURCE</th><th>IDENTIFIER</th></tr></thead><tbody><tr><td colspan="3">Antibodies</td></tr><tr><td>Chicken polyclonal anti-GFP (1/1000)</td><td>Sigma-Aldrich</td><td>Cat#AB16901,<br>RRID:AB_90890</td></tr><tr><td>Alexa Fluor™ 488-conjugated anti-chicken IgY (H+L) (1/1000)</td><td>Invitrogen</td><td>Cat#A-11039,<br>RRID:AB_2534096</td></tr><tr><td colspan="3">Bacterial and virus strains</td></tr><tr><td>pIRES2-CMV-nAChRb2(E61C)-IRES-GFP</td><td>Durand-de Cuttoli et al.<sup>13</sup>, Addgene</td><td>Cat#164779,<br>RRID:Addgene_164779</td></tr><tr><td>pLenti-pGK-nAChRb2(E61C)-IRES-GFP</td><td>Durand-de Cuttoli et al.<sup>13</sup>, Addgene</td><td>Cat#164777,<br>RRID:Addgene_164777</td></tr><tr><td>AAV5-GFP: ssAAV5/2-hSyn1-EGFP-WPRE-hGHp(A)</td><td>ETH Zürich Viral Vector Facility</td><td>Cat#v81-5</td></tr><tr><td colspan="3">Chemicals, peptides, and recombinant proteins</td></tr><tr><td>MPEG<sub>4</sub>Ch chloride</td><td>Jehl et al.<sup>1</sup>, custom-synthesized by Broadpharm®</td><td>Cat#BP-24475,<br>PubChem CID 146020404</td></tr><tr><td>MPEG<sub>4</sub>Ch Stock solution</td><td>custom prep, see Materials and equipment setup</td><td>N/A</td></tr><tr><td>Dimethyl sulfoxide (DMSO)</td><td>Fisher Scientific</td><td>Cat#D12345</td></tr><tr><td>Sodium Chloride (NaCl)</td><td>Sigma-Aldrich</td><td>Cat#S7653</td></tr><tr><td>Nicotine hydrogen tartrate salt</td><td>Glentham LifeSciences</td><td>Cat# GL9693</td></tr><tr><td>Sodium hydroxide (NaOH)</td><td>Sigma-Aldrich</td><td>Cat#S5881</td></tr><tr><td>Potassium chloride (KCl)</td><td>Sigma-Aldrich</td><td>Cat#P9333</td></tr><tr><td>Monosodium phosphate (NaH<sub>2</sub>PO<sub>4</sub>)</td><td>Sigma-Aldrich</td><td>Cat#S8282</td></tr><tr><td>Calcium chloride (CaCl<sub>2</sub>)</td><td>Sigma-Aldrich</td><td>Cat#233506</td></tr><tr><td>Magnesium chloride (MgCl<sub>2</sub>)</td><td>Sigma-Aldrich</td><td>Cat#M2670</td></tr><tr><td>Sodium Bicarbonate (NaHCO<sub>3</sub>)</td><td>Sigma-Aldrich</td><td>Cat#S6297</td></tr><tr><td>Glucose</td><td>Sigma-Aldrich</td><td>Cat#S0389</td></tr><tr><td>Sucrose</td><td>Sigma-Aldrich</td><td>Cat#49159</td></tr><tr><td>Kynurenic acid</td><td>Sigma-Aldrich</td><td>Cat#K3375</td></tr><tr><td>K-gluconate</td><td>Sigma-Aldrich</td><td>Cat#P1847</td></tr><tr><td>HEPES</td><td>Sigma-Aldrich</td><td>Cat#H3375</td></tr></tbody></table>
<table><tbody><tr><td>Ethylene glycol tetraacetic acid (EGTA)</td><td>Sigma-Aldrich</td><td>Cat#E3889</td></tr><tr><td>ATP magnesium salt (Mg-ATP)</td><td>Sigma-Aldrich</td><td>Cat#A9187</td></tr><tr><td>K-gluconate solution</td><td>custom prep, see Materials and equipment setup</td><td>N/A</td></tr><tr><td>1x aCSF solution</td><td>custom prep, see Materials and equipment setup</td><td>N/A</td></tr><tr><td>Sucrose solution</td><td>custom prep, see Materials and equipment setup</td><td>N/A</td></tr><tr><td>Ketamine (Imalgene 1000)</td><td>Merial</td><td>N/A</td></tr><tr><td>Xylazine (Rompun 2%)</td><td>Bayer</td><td>N/A</td></tr><tr><td>Heparin</td><td>Sigma-Aldrich</td><td>Cat#PHR8927</td></tr><tr><td>Chloral hydrate</td><td>Sigma-Aldrich</td><td>Cat#302-17-0</td></tr><tr><td>Isoflurane</td><td>Piramal Healthcare</td><td>N/A</td></tr><tr><td>Vaseline</td><td>Vaseline</td><td>N/A</td></tr><tr><td>Mineral oil</td><td>Carlo Erba</td><td>Cat#356601</td></tr><tr><td>Vybrant™ Multicolor Cell-Labeling Kit (DiO, Dil, DiD Solutions, 1 mL each)</td><td>Invitrogen</td><td>Cat#V22889</td></tr><tr><td>PBS 10X</td><td>Sigma-Aldrich</td><td>Cat#70011044</td></tr><tr><td>4% Paraformaldehyde (PFA)</td><td>Thermo Scientific</td><td>Cat#J19943.K2</td></tr><tr><td colspan="3">Experimental models: Organisms/strains</td></tr><tr><td>Mouse: C57BL/6NRj (males and females, 8–12 weeks)</td><td>Janvier Labs</td><td>RRID:IMSR_RJ:C57 BL-6JRJ</td></tr><tr><td>Mouse: Chrnb4<sup>E61C</sup> (males and females, 8–12 weeks)</td><td>Jehl et al.<sup>1</sup>, provided by Phenomin</td><td>N/A</td></tr><tr><td colspan="3">Software and algorithms</td></tr><tr><td>Clampfit and Clampex (part of pClamp 10.6 suite)</td><td>Molecular Devices</td><td>RRID:SCR_011323</td></tr><tr><td>Cheetah software</td><td>Neuralynx</td><td>Cat#36-0301-0054, https://neuralynx.fh-co.com/research-software/cheetah/</td></tr><tr><td>CSC Spike Extractor software</td><td>Neuralynx</td><td>N/A</td></tr><tr><td>SpikeSort3D software</td><td>Neuralynx</td><td>https://neuralynx.fh-co.com/research-software/spikesort-3d/</td></tr><tr><td>R Statistical Software</td><td>https://www.r-project.org</td><td>RRID:SCR_001905</td></tr><tr><td>ezTrack</td><td>Pennington et al.<sup>33</sup></td><td>RRID:SCR_021496</td></tr><tr><td>FIJI</td><td>Schindelin et al.<sup>34</sup></td><td>RRID:SCR_002285</td></tr><tr><td>DMC-BrainMap</td><td>Jung et al.<sup>35</sup></td><td>https://github.com/hejDMC/napari-dmc-brainmap</td></tr><tr><td colspan="3">Other</td></tr><tr><td>QuikChange II XL Site-Directed Mutagenesis Kit</td><td>Agilent</td><td>Cat#200522</td></tr><tr><td>Practum laboratory balance, model 224</td><td>Sartorius</td><td>Cat#practum224-1S</td></tr><tr><td>Spoon spatula</td><td>Dutscher</td><td>Cat#3340</td></tr><tr><td>PIPETMAN® P10 micropipette</td><td>Gilson</td><td>Cat#F144802</td></tr></tbody></table>
<table><tr><td>PIPETMAN® P1000 micropipette</td><td>Gilson</td><td>Cat#F144059M</td></tr><tr><td>Microcentrifuge tube, 1.5 mL</td><td>Eppendorf</td><td>Cat#0030108116</td></tr><tr><td>Microtube, 0.2 mL</td><td>Eppendorf</td><td>Cat#0030124332</td></tr><tr><td>Microtube, 0.5 mL</td><td>Eppendorf</td><td>Cat#0030123603</td></tr><tr><td>Laboratory bottle, 1 L</td><td>Schott Duran</td><td>Cat#218015455</td></tr><tr><td>Vortex Genie 2</td><td>Scientific industries</td><td>Cat#SI-0256</td></tr><tr><td>25 mm sterile nylon membrane, 0.2µm pore size</td><td>Nalgene</td><td>Cat#726-2520</td></tr><tr><td>Bottle-top sterile filter, 0.2 µm pore size</td><td>Nalgene</td><td>Cat#Z358223</td></tr><tr><td>Magnetic hotplate stirrer</td><td>LMS</td><td>Cat#hts-1003</td></tr><tr><td>Magnetic stir bar</td><td>Fisherbrand</td><td>Cat#001.3001</td></tr><tr><td>SevenEasy pH meter</td><td>Mettler-Toledo</td><td>Cat#S20</td></tr><tr><td>Sonicator</td><td>BPAC</td><td>Cat#bpac03sk120s</td></tr><tr><td>Parafilm tape M</td><td>Amcor</td><td>Cat#9170002</td></tr><tr><td>Luer Lock Syringe, 50 mL</td><td>Terumo</td><td>Cat#SS+50L1</td></tr><tr><td>Peristaltic pump Ismatec IPC-N-08</td><td>Ismatec</td><td>Cat#ISM936D</td></tr><tr><td>Compresstome VF-200</td><td>Precisionary Instruments</td><td>Cat#VF-200</td></tr><tr><td>Costar® 12-well Clear TC-treated Multiple Well Plates</td><td>Corning</td><td>Cat#3513</td></tr><tr><td>P-87 Flaming/Brown Micropipette Puller</td><td>Sutter Instruments</td><td>Cat#P87</td></tr><tr><td>Thin-wall borosilicate glass</td><td>Warner Instruments</td><td>Cat#G150TF-3</td></tr><tr><td>Microloader</td><td>Eppendorf</td><td>Cat#EP5242956003</td></tr><tr><td>Axoclamp 200B Microelectrode Amplifier</td><td>Molecular Devices</td><td>Cat#AxOPATCH 200B-2</td></tr><tr><td>Digidata 1550A Low-Noise Data Acquisition Digitizer</td><td>Molecular Devices</td><td>Cat#5035919A</td></tr><tr><td>Micromanipulator</td><td>Scientifica</td><td>Cat#PS-7550</td></tr><tr><td>Picopritzer Pneumatic Pump</td><td>World Precision Instruments</td><td>Cat#PV800</td></tr><tr><td>GeneClamp 500 amplifier</td><td>Axon Instruments</td><td>N/A</td></tr><tr><td>8-way, 12-way, and 2-way 4-port electric rotary valve</td><td>Bio-Chem Fluidics</td><td>https://www.arcmedgroup.com/pumps-valves#electric-rotary-valves</td></tr><tr><td>SliceScope upright microscope</td><td>Scientifica</td><td>Cat#S-Scope-II</td></tr><tr><td>ROLERA™ Bolt Scientific CMOS Camera</td><td>QImaging</td><td>Cat#01-ROL-BOLT-M-12</td></tr><tr><td>Halogen lamp unit</td><td>Olympus</td><td>Cat#TH4-200</td></tr><tr><td>PIPETMAN® P2 micropipette</td><td>Gilson</td><td>Cat#F144801</td></tr><tr><td>Gas anesthesia station for small animals</td><td>TEMSEGA</td><td>Cat#MiniHUB-V3</td></tr><tr><td>Charcoal anesthesia gas filter</td><td>F/AIR</td><td>Cat#80120</td></tr><tr><td>Dual Small Animal Stereotaxic w/ Digital Display</td><td>David Kopf Instruments</td><td>Cat#942-WA</td></tr><tr><td>ThermoStar Homeothermic System</td><td>RWD</td><td>Cat#69027</td></tr><tr><td>Routine stereo microscope</td><td>Leica Biosystems</td><td>Cat#M80</td></tr><tr><td>Drilling machine</td><td>Foredom</td><td>Cat# K.1070</td></tr><tr><td>0.7 mm steel drill burrs</td><td>Fine Science Tools</td><td>Cat#19009-07</td></tr><tr><td>Isis rodent shaver</td><td>Aesculap</td><td>Cat#GT421</td></tr><tr><td>100 µL, Gastight Syringe, Small Removable Needle</td><td>Hamilton</td><td>Cat#1710</td></tr><tr><td>Polyethylene Tubing</td><td>Instech</td><td>Cat#BTPE-10</td></tr><tr><td>Loctite Superglue-3 original</td><td>Henkel</td><td>Cat#2959596</td></tr></table>
<table><tr><td>ToughCut fine scissors</td><td>Fine Science Tools</td><td>Cat#14058-09</td></tr><tr><td>Dumont #5 forceps</td><td>Fine Science Tools</td><td>Cat#11251-20</td></tr><tr><td>Olsen-Hegar Needle Holders with Suture Cutters</td><td>Fine Science Tools</td><td>Cat#12002-12</td></tr><tr><td>Instrument Sterilization Container</td><td>Fine Science Tools</td><td>Cat#20840-02</td></tr><tr><td>Autoclave</td><td>Matachana</td><td>Cat#S1000</td></tr><tr><td>Thomas Tetrode Mini Matrix microdrive system</td><td>Thomas Recording</td><td>Cat#AN000423</td></tr><tr><td>Tetrode for Tetrode Mini Matrix (Tip A)</td><td>Thomas Recording</td><td>Cat#AN000259</td></tr><tr><td>Quartzglass microinjection pipettes for Thomas microdrive</td><td>Thomas Recording</td><td>Cat#AN000155</td></tr><tr><td>Digital Lynx 4SX</td><td>Neuralynx</td><td>Cat#31-0605-0141, https://neuralynx.fh-co.com/research-hardware/data-acquisition/digital-lynx-sx/</td></tr><tr><td>PC-10 puller</td><td>Narishige</td><td>RRID:SCR_022057</td></tr><tr><td>Capillary micropipette for PCR 10 µL</td><td>Drummond</td><td>Cat#5-000-1001-X10</td></tr><tr><td>One-axis Oil Hydraulic Micromanipulator</td><td>Narishige</td><td>Cat#MO-10</td></tr><tr><td>Y-maze apparatus</td><td>Imetronic</td><td>https://www.imetroni-c.com/devices/y-maze/</td></tr><tr><td>Intel RealSense Camera</td><td>Intel</td><td>Cat#D455</td></tr><tr><td>Surfa Safe™ Premium disinfectant spray</td><td>Laboratoires Anios</td><td>Cat#2419326</td></tr><tr><td>Vibratome VT1000 S</td><td>Leica Biosystems</td><td>RRID:SCR_016495</td></tr></table>

## Materials and equipment setup

### Preparation of MPEG₄Ch stock solution

* To prepare 0.5 mL of a 50 mM stock solution of MPEG₄Ch retrieve MPEG₄Ch chloride (MW = 394.9 g/mol) powder stored at -20°C.

**Note:** As a general rule, chemicals should be stored at the highest practical concentration (Here 50 mM). However, a lower concentration (e.g. 10 mM) is also acceptable.

**Note:** Salts—such as the chloride salt added to MPEG₄Ch in this case—are generally more soluble in water than their uncharged counterparts. This increased solubility makes them preferable for *in vivo* use.

* Weigh 9.87 mg of MPEG₄Ch chloride using a clean, dry spatula.

* Transfer the powder into a sterile microcentrifuge tube.

* Just before use, open a fresh ampoule of anhydrous dimethyl sulfoxide (DMSO).

**CRITICAL:** DMSO must be strictly anhydrous. Maleimide groups in MPEG₄Ch react with water, compromising compound integrity. Do not use previously opened DMSO, even if tightly capped, as it may have absorbed moisture from the air.

* Add 500 µL of the freshly opened anhydrous DMSO to the tube containing MPEG₄Ch.
* Mix gently by pipetting up and down or vortexing until fully dissolved.

* Aliquot 10 µL portions into clearly labeled 0.2 mL Eppendorf tubes.

**Note:** Store aliquots at -20°C and minimize air exposure by using one aliquot per experiment and keeping tubes sealed. Avoid repeated freeze-thaw cycles.

## Preparation of filtered 0.9% saline solution

* Add 9 g of Sodium Chloride (NaCl; Sigma-Aldrich S7653) and 950 mL of milli-Q water to a 1 L bottle.

* Stir at medium speed until fully dissolved and adjust the volume up to 1 L with milli-Q water.

* Filter the solution using a 50 mL syringe (Terumo Luer Lock Syringe, 50 mL) and an appropriate syringe filter, such as Nalgene 25 mm syringe filter (Sterile Nylon membrane, 0.2µm pore size).

**Note:** Store the bottle at 4°C (up to one year – date the bottle).

## Preparation of nicotine tartrate solution (57 mg/L):

* Pour 100 mL of sterile, filtered 0.9% saline into a clean beaker.

* Weigh and add 5.7 mg of nicotine hydrogen tartrate salt (Glentham LifeSciences, GL9693, MW = 462.41 g/mol) to the saline, yielding a final concentration of 57 mg/L.

* Stir the solution thoroughly until the salt is completely dissolved.

* Adjust the pH to 7.2 ± 0.2 using sodium hydroxide (NaOH) as needed.

**Note:** This solution is designed to deliver a dose of 0.2 mg/kg nicotine free base per injection, using an injection volume of 0.1 mL per 10 g body weight (for a 25 g mouse, inject 0.25 mL). This corresponds to an injection volume of 10 mL/kg.

**Note:** Prepare the 57 mg/L solution fresh during the week of the experiment, or alternatively, prepare a more concentrated stock solution, aliquot, and store at -20 °C. Thaw individual aliquots and dilute to the working concentration as needed.

## Preparation of artificial cerebrospinal fluid (aCSF) 10x stock solution

<table><thead><tr><td>Reagent</td><td>Final Concentration (mM)</td><td>Amount</td></tr></thead><tbody><tr><td>milli-Q water</td><td>-</td><td>1 L</td></tr><tr><td>NaCl</td><td>1250</td><td>73.05 g</td></tr><tr><td>KCl</td><td>25</td><td>1.86 g</td></tr><tr><td>NaH<sub>2</sub>PO<sub>4</sub></td><td>12.5</td><td>1.5 g</td></tr><tr><td>NaHCO<sub>3</sub></td><td>260</td><td>21.85 g</td></tr></tbody></table>

**Note:** Place a clean 1 L beaker with a magnetic stir bar on a stirring plate and add 500 mL of milli-Q water. While stirring, add NaCl, KCl, NaH<sub>2</sub>PO<sub>4</sub>, and NaHCO<sub>3</sub>. Continue stirring until all components are fully dissolved. Once everything is dissolved, transfer the solution into a clean 1 L volumetric flask and fill up to the 1 L mark with milli-Q water. Place a 0.2 µm membrane bottle-top filter (Nalgene) on a clean 1 L collection bottle, connect it to a vacuum source, and pour the solution into the filter reservoir. Allow
283 the solution to fully filter into the bottle. Label the final solution appropriately and store as needed.

284 Store the final solution at 4°C and use within 30 days.

285

## Preparation of aCSF 1x

<table><thead><tr><td>Reagent</td><td>Final Concentration (mM)</td><td>Amount</td></tr></thead><tbody><tr><td>milli-Q water</td><td>N/A</td><td>1 L</td></tr><tr><td>CaCl<sub>2</sub> 2H<sub>2</sub>O 2 M</td><td>2</td><td>1 mL</td></tr><tr><td>MgCl<sub>2</sub> 6H<sub>2</sub>O 2 M</td><td>1</td><td>0.5 mL</td></tr><tr><td>Glucose</td><td>10</td><td>1.98 g</td></tr><tr><td>Sucrose</td><td>15</td><td>5.13 g</td></tr><tr><td>aCSF 10x stock solution</td><td>N/A</td><td>100 mL</td></tr></tbody></table>

288

289 **Note:** Place a clean 1 L beaker on a magnetic stirring plate and add a clean magnetic stir bar. Pour 500

290 mL of milli-Q water into the beaker. While stirring, add the CaCl₂ and MgCl₂ stock solutions, then weigh

291 and add glucose and sucrose. Stir until fully dissolved. Using a volumetric flask, measure and add the

292 aCSF 10x stock solution to the beaker. Rinse the volumetric flask with a small amount of milli-Q water

293 and pour the rinse into the beaker to recover any remaining aCSF 10x stock solution. Transfer the

294 contents of the beaker into a clean 1 L volumetric flask and fill up to the 1 L mark with milli-Q water.

295 Mix by gently inverting the volumetric flask several times, then transfer the final solution into a clean 1

296 L bottle for storage. Store the final solution at 4°C and use within 3 days.

297

## Preparation of sucrose-based cutting solution

<table><thead><tr><td>Reagent</td><td>Final Concentration (mM)</td><td>Amount</td></tr></thead><tbody><tr><td>milli-Q water</td><td>N/A</td><td>1 L</td></tr><tr><td>MgCl<sub>2</sub> 6H<sub>2</sub>O 2 M</td><td>5.9</td><td>2.95 mL</td></tr><tr><td>Glucose</td><td>2.5</td><td>0.49 g</td></tr><tr><td>Sucrose</td><td>25</td><td>8.55 g</td></tr><tr><td>aCSF 10x stock solution</td><td>N/A</td><td>100 mL</td></tr><tr><td>Kynurenic Acid</td><td>N/A</td><td>200 mg</td></tr></tbody></table>

300

301 **Note:** Place a clean 1 L beaker on a magnetic stirring plate and add a clean magnetic stir bar. Pour 500

302 mL of milli-Q water into the beaker. While stirring, add the MgCl₂ stock solution. Weigh and add the

303 glucose and sucrose. Using a 100 mL volumetric flask, measure and add 100 mL of aCSF 10x stock

304 solution to the beaker. Rinse the volumetric flask with a small amount of milli-Q water and pour the

305 rinse into the beaker to recover any remaining 10x aCSF. Transfer the contents of the beaker into a

306 clean 1 L volumetric flask and fill to the 1 L mark with milli-Q water. Mix by gently inverting the

307 volumetric flask several times, then transfer the solution into a clean 1 L storage bottle. Weigh 200 mg

308 of kynurenic acid and add it to the solution. Sonicate for 15 minutes to ensure complete dissolution.

309 Store the final solution at 4°C and use within 3 days.

310

## Preparation of K-gluconate internal solution

311
<table><thead><tr><td>Reagent</td><td>Final Concentration (mM)</td><td>Amount</td></tr></thead><tbody><tr><td>milli-Q water</td><td>N/A</td><td>50 mL</td></tr><tr><td>K-gluconate</td><td>135</td><td>1.5812 g</td></tr><tr><td>HEPES (free acid)</td><td>10</td><td>119.15 mg</td></tr><tr><td>KCl</td><td>5</td><td>18.6375 mg</td></tr><tr><td>MgCl<sub>2</sub></td><td>2</td><td>20.33 mg</td></tr><tr><td>EGTA</td><td>0.1</td><td>1.9018 mg</td></tr><tr><td>Mg-ATP</td><td>2</td><td>50.718 mg</td></tr></tbody></table>

**Note:** Place a clean 50 mL glass beaker on a magnetic stir plate and add a magnetic stir bar. Pour 35 mL of milli-Q water into the beaker and begin stirring. While stirring, weigh and add the following components in sequence: K-gluconate, HEPES (free acid), KCl, MgCl₂, EGTA, and Mg-ATP. Stir until all solids are fully dissolved. Adjust the pH to 7.2 using small volumes of HCl (e.g., 0.1–1 M), monitoring with a calibrated pH meter. Transfer the solution to a clean 50 mL volumetric flask and bring the final volume up to 50 mL with milli-Q water. Seal the flask with Parafilm and mix gently by inverting several times. The solution should then be filter-sterilized using a 0.2 µm pore syringe filter (Nalgene) into a sterile container. Aliquot 0.5 mL of the internal solution into sterile, labeled 0.5 mL Eppendorf tubes. Store aliquots at –20 °C and use within 6 months. Use thawed aliquots on the day of the experiment. Do not refreeze. Discard any remaining solution after the recording session to avoid degradation or contamination.

## Preparation of choral hydrate solution for anesthesia

* To prepare 100 mL of an 8% (w/v) solution, weigh out 8 grams of chloral hydrate powder (Sigma-Aldrich 15307).

* Add the powder to 80 mL of milli-Q water in a beaker or flask.

* Stir the solution using a magnetic stir bar until the powder is completely dissolved.

* Transfer the solution to a graduated cylinder or volumetric flask and add milli-Q water to bring the final volume up to 100 mL.

* Filter the solution through a 0.22 µm sterile syringe filter into a sterile container. Store at 4°C and protect from light.

**Note:** Chloral hydrate is hazardous and should be handled with appropriate PPE. Safer alternatives, such as isoflurane, may be used if preferred or required by institutional guidelines.

## Surgical setup for in vivo electrophysiology and MPEG₄Ch injection

Timing: 30 min

* Surgical Instruments and supplies

* Pulled glass micropipettes (Drummond 10 µL PCR Pipettes): For microinjections.

* Microinjector (e.g., Narishige MO-10 or IMS-30)

* P2 micropipette: For precise liquid handling.

* Parafilm: For preparing and handling small volumes of solution to infuse.

* Mineral oil: For filling micropipettes and preventing evaporation.

* Vaseline: For lubricating pipette plungers.
350

351

352

353

354

355

356

357

358

359

360

361

362

363

364

365

### • Anesthesia and Fluids

366

367

368

369

370

371

372

373

374

375

376

377

378

379

380

381

382

383

384

385

386

387

388

389

### • Sterilization and Storage

390

391

392

393

* Shaver: For removing fur from the surgical area.

* Heating pad/mat (set to 37°C): To maintain animal body temperature during surgery.

* Stereotaxic frame (e.g., Kopf) with stereotaxic arm, ear bars and gas mask: For head fixation and anesthesia delivery.

* Stereo microscope (e.g. Leica M80): for precise visualization

* Fine scissors and forceps (e.g., FST instruments): For precise tissue handling and dissection.

* Drilling machine with 0.9 mm steel drill bit: For craniotomy.

* Sutures thread with needle and needle holders: For wound closure.

* Cotton buds: For cleaning and hemostasis.

* Betadine: For skin disinfection.

* Physiologic gel: To protect eyes from drying during anesthesia.

* Sterile gloves: For aseptic technique.

* Isoflurane: For inhalation anesthesia (3-4% for induction and 1-2% for maintenance).

* Oxygen source: For delivering anesthetic gases.

* Ketamine (100 mg/mL): Used for injectable anesthesia, typically combined with xylazine.

* Xylazine (Rompun®, 20 mg/mL): Administered in combination with ketamine for deep surgical anesthesia.

* 0.9% Saline: For hydration and solution preparation.

* Lidocaine: For local anesthesia at incision site. Prepare a solution at 1 mg/mL (by diluting 10 µL of 10 mg/mL stock in 90 µL saline). Inject 0.125 mL per animal, corresponding to a dose of 5 mg/kg (for a 25 g mouse, this is 0.005 mL/g).

* Buprenorphine: For perioperative analgesia. Prepare a solution at 0.03 mg/mL (by diluting 20 µL of stock solution in 180 µL of physiological saline). Inject 0.05 mL (50 µL) subcutaneously per mouse, corresponding to a dose of 0.06 mg/kg (for a 25 g mouse, this is 0.002 mL/g).

* MPEG₄Ch stock solution (700nl, 50 mM in water-free DMSO, -20°C): For CATCH application.

* AAV5-GFP: For injection site verification via fluorescence. AAV5-GFP refers to the ssAAV5/2-hSyn1-EGFP-WPRE-hGHp(A) viral vector (see Resources Table). The viral stock used had a titer of 5.7 × 10¹¹ vg/mL.

* Fluorescent dyes (Dil, DiO, DiD; 1 mg/mL): For visualizing injection sites.

* Autoclave: For sterilizing surgical instruments.

* Sterile storage containers: For keeping tools and reagents contamination-free.

* Charcoal anesthesia gas filter: For isoflurane waste gas scavenging.
<figure><img src="image_3.png" /></figure>

394 **Note:** Prepare all instruments and reagents before beginning surgery to minimize anesthesia time.
395 Sterilize surgical instruments by autoclaving them beforehand at 121°C for 30 min. Maintain aseptic
396 technique throughout all surgical procedures. Ensure all solutions are freshly prepared and filtered as
397 described in the protocol.

398 Step-by-step method details

399
400 *Ex vivo electrophysiology recording after incubation with MPEG4Ch*
401
402 This procedure describes how to assess the efficacy and specificity of CATCH-mediated receptor
403 antagonism at the cellular level using acute brain slices. As an example, this approach was employed in
404 Jehl et al.¹ to quantify the impact of MPEG4Ch incubation on nAChR function in the interpeduncular
405 nucleus (IPN) of Chrnb4E61C KI mice.
406

407 **Note:** By comparing nicotine-evoked currents in slices treated with MPEG₄Ch versus untreated controls, this method provides a direct and quantitative readout of receptor blockade and functional receptor expression levels.
408
409
410

411 **CRITICAL:** This *ex vivo* validation is critical for confirming the molecular and functional specificity of
412 CATCH prior to *in vivo* or behavioral applications.
413

414 Timing: 3h
415

416
417 1. Dissection and slice preparation.³⁶ Timing: 20-30min.
418 a) Prepare the dissection setup.

419 **CRITICAL:** Ensure all solutions (sucrose-based cutting solution, 1x aCSF) are freshly prepared in the
420 morning, stored appropriately (max 3 days at 4°C), and fully saturated with carbogen gas (95% O₂ / 5%
421 CO₂).

422 b) Chill the cutting solution to ≈0°C in the freezer.

423 **Note:** Keep the cutting solution on ice throughout the dissection and slicing steps. The detailed
424 procedure for preparing these solutions can be found in the *Materials and Equipment Setup* section of
425 this protocol.

426 c) Anesthetize a KI mouse via intraperitoneal injection of ketamine/xylazine (0.11 mL
427 ketamine [100 mg/mL] + 0.06 mL xylazine [20 mg/mL] in 0.63 mL sterile NaCl). Confirm
428 anesthesia via loss of pedal reflex before proceeding.
429

429 d) Perform transcardiac perfusion:

430 i) Open the thoracic cavity.

431 ii) Inject 0.1 mL of heparin (1000 U/mL) into the apex of the heart.

432 iii) Clamp the descending aorta
<figure><img src="image_4.png" /></figure>

433 iv) perfuse with ice-cold, oxygenated sucrose-based cutting solution using a
434 peristaltic pump (Ismatec IPC) for 1 min 20 sec.

435 e) Decapitate the mouse and extract the brain.

436 f) Immediately submerge the brain in 0°C oxygenated cutting solution.

437 g) Place the brain ventral side down on filter paper and make two coronal cuts: one
438 behind the olfactory bulbs and one through the cerebellum.

439
440 **Note:** This protocol targets the IPN; adjust cutting if targeting other regions. IPN coordinates can range
441 between Antero-Posterior (AP) -3.3 to -3.6 mm; Medio-lateral (ML) 0.35-0.5 mm; Dorso-ventral (DV)
442 4.2-4.9 mm, from Bregma.

443
444 h) Affix the rostral brain section onto the specimen holder of a Compresstome using glue
445 and supportive agarose gel.

446 i) Fill the slicing chamber with 150 mL ice-cold oxygenated cutting solution.

447 j) Slice the brain into 250 µm coronal sections using the Compresstome.

448 k) Transfer slices containing the IPN into a 35°C oxygenated sucrose-based recovery
449 solution for 8 minutes. Repeat for additional IPN slices (typically 2-3 slices per brain).

450 l) Move slices to oxygenated 1x aCSF (kept at 20-25°C).

451 i) Incubate for at least 60 minutes.

452 m) Clean all dissection instruments thoroughly with distilled water and, when
453 appropriate, 70% ethanol.

454
455 2. MPEG₄Ch incubation (Figure 2A). Timing: 20min.

456 **Note:** This protocol is optimized for between-subject comparisons. Ideally, control and MPEG4Ch-
457 treated slices should be recorded on the same day, with control recordings performed first, because a
458 slice cannot serve as its own control after MPEG4Ch incubation. However, in practice, we found that
459 slice health can decline over the course of a recording session, potentially confounding the results. To
460 minimize this issue, we performed control and treated recordings on separate days, ensuring that each
461 slice was in optimal condition at the time of recording.

462 **Note:** As an alternative to in-slice MPEG₄Ch incubation, MPEG₄Ch can be locally injected in the IPN *in vivo* in KI mice (Figure 2B). Patch-clamp recordings can then be performed on acute brain slices without further incubation, since the compound is already present in the tissue (see step 17). Our *ex vivo* data show that MPEG₄Ch-mediated antagonism of β4 nAChR subunits is detectable as early as 1 day after infusion and lasts for at least 10 days;¹ longer durations should be validated experimentally.

467 a) Freshly prepare 700 µL of MPEG₄Ch-containing 1x aCSF by diluting 0.7 µL of 50 mM
468 MPEG₄Ch stock (in DMSO) to achieve 50 µM final concentration.

469 **CRITICAL:** Prepare the 50 µM MPEG₄Ch solution fresh on the day of the experiment, since maleimide
470 groups in MPEG₄Ch react with water, compromising compound integrity. Store the MPEG₄Ch stock
471 solution (50 mM in DMSO) at -20°C. Avoid repeated freeze-thaw cycles by aliquoting the stock
472 solution as needed.
474 **Note:** Lower concentrations, such as 10 µM, should also be effective, but concentrations higher than 475 50 µM are not recommended for slice experiments.
476 b) After the recovery period, transfer the slice containing the IPN to a small incubation 477 chamber (using a 12 well plates) filled with oxygenated 1x aCSF, continuously 478 bubbled with 95% O₂ / 5% CO₂.
479 c) Remove the existing 1x aCSF with a transfer pipette and immediately replace it with 480 the 700 µL of MPEG₄Ch-containing 1x aCSF, ensuring continuous oxygenation.
481 d) Incubate the slice for 10 minutes under constant oxygenation.
482 e) **Rinsing:** Remove the MPEG₄Ch-containing solution and replace it with 700 µL of fresh 483 oxygenated 1x aCSF. Allow the slice to rest for 3 minutes.
484 i) Repeat this rinsing step two more times for a total of three rinses.
485 f) Transfer the slice to the patch clamp recording chamber.
486
487 **Note:** Handle the slice with extreme care to prevent damage during transfers and throughout the 488 oxygenation process.

489 3. Patch-clamp electrophysiology: Pre-recording preparation. Timing: 15min.
490 a) **Pipette preparation:** Pull patch pipettes (3–6 MΩ) from borosilicate glass capillaries
491 (G150TF-3, Warner Instruments) using a micropipette puller (P-87, Sutter
492 Instruments, Novato, CA).

493 **Note:** This resistance range is suitable for whole-cell recordings in most neurons, though slight
494 adjustments may be required based on cell type and morphology.

495 b) Thaw an aliquot of K-gluconate internal solution (preparation detailed in *Materials and Equipment setup* section).
496 i) To fill pipettes with intracellular solution, draw the aliquot into a 1 mL syringe
497 fitted with a 4 µm Nalgene syringe filter and a microloader (Eppendorf).
498 ii) Fill pipettes until the solution reaches the electrode tip, then gently flick the
499 pipette to remove any air bubbles.
500

501 **Note:** Use the same method to fill puff pipettes with nicotine-containing 1x aCSF (30 µM), but omit
502 the syringe filter.

503 c) **Perfusion and positioning:**
504 i) Power on the patch clamp rig (Axopatch 200B amplifier connected to a
505 Digidata 1550 Low-Noise acquisition system, Molecular Devices, USA) and acquisition
506 software (Clampex software, pClamp 10.5, Molecular Devices).
507 ii) Begin perfusion of fresh oxygenated 1x aCSF into the recording chamber at a
508 flow rate of 0.5–1 mL/min via gravity feed or peristaltic pump.
509 iii) Direct the waste line into a 2 L Erlenmeyer flask containing diluted bleach,
510 connected to a vacuum line.
511 d) Once perfusion is stable, use a transfer pipette to gently place the slice in the
512 recording chamber.
513 e) Secure the slice with a stainless-steel slice anchor (harp) fitted with nylon threads.
514 f) Mount the patch and puff pipettes onto their respective headstages.

515 g) Check slice viability, [Troubleshooting 1](#).

516

517 4. Patch-clamp electrophysiology: Patching. Timing: 3-4h.

518 a) Apply positive pressure to the patch pipette using a syringe.

519 b) Lower both pipettes into the recording bath.

520 c) Confirm the resistance of the recording pipette is 3–6 MΩ.

521 d) Bring both pipettes above the selected neuron using micromanipulators (Scientifica) and zero the pipette offset using the “membrane test” function on the Clampex software.

524 e) Slowly descend the patch pipette until a small dimple and increased resistance are observed.

526 f) Release the positive pressure and look for a further resistance increase.

527 g) Gently apply suction via the syringe while holding the potential at –40 mV to form a giga-ohm seal.

529 h) Once the seal is stable, hold the membrane potential at –70 mV and apply brief, gentle suction to break into whole-cell configuration.

531

532 **Note:** Monitor the capacitive transient and adjust compensation on the amplifier as needed when breaking into whole-cell mode.

534

535 i) Set the amplifier to **voltage-clamp mode** and begin recording using a **gap-free** protocol.

537 j) Position the puff pipette ~20 µm from the patched cell and adjust its height to match the patch pipette.

539 k) Use a picopritzer pneumatic pump (PV800, WPI) connected to a pressurized air system to deliver puffs of nicotine (30 µM) at a starting pressure of 5–10 psi.

541 **Note:** Start with minimal puff pressure and adjust carefully to avoid disturbing the patched cell. The first puff typically delivers a lower concentration of nicotine due to aCSF diffusion into the pipette; therefore, 2–3 puffs may be needed to ensure stable and reliable responses. [Troubleshooting 2](#).

544 l) Following recordings, clean all tools, tubing, and perfusion lines with milli-Q water and, where appropriate, 70% ethanol.

546

547 5. Data export and analysis

548 a) Export raw current traces from Clampfit or compatible acquisition software.

549 b) Analyze peak nicotine-induced current amplitudes in R or Python, depending on the experimental goal. [Troubleshooting 3](#).

551 c) **Example Goal:** Compare peak current amplitudes in IPN neurons between MPEG₄Ch-treated and untreated (control) slices in KI mice.

553 **Note:** The statistical method and sample size depend on the specific hypothesis. For comparisons, use t-tests or linear mixed-effects models as appropriate.

554
In vivo acute electrophysiology recording after local infusion of MPEG₄Ch

This section describes how to apply the CATCH protocol to investigate the functional consequences of localized receptor antagonism on neuronal spiking activity. It describes the recording and analysis of neuronal activity in the IPN of anesthetized mice following local infusion of MPEG₄Ch and subsequent nicotine injections, using tetrode-based electrophysiology.

Note: This protocol can be adapted for use in other brain regions and with different receptor types.

Timing: 1 day

6. Prior to beginning the procedure gather all the supplies listed in the *Materials and equipment setup* section.

7. Prepare a fresh dilution of MPEG₄Ch solution for *in vivo* use. Timing: 5 min
   a) Take MPEG₄Ch stock solution (50mM) stored at -20°C in water-free DMSO.
   b) Prepare 0.9% Saline Solution as described in *Materials and equipment setup* section.
   c) Dilute MPEG₄Ch Stock Solution.
   i) In a 1 mL Eppendorf tube, add 100 µL of filtered saline and 0.5 µL of MPEG₄Ch stock solution (50mM) to achieve a final concentration of 250 µM for *in vivo* use.

**CRITICAL:** Prepared MPEG₄Ch solution should be used within the same day. Prepare a fresh dilution daily before use.

Note: Ensure solution is at the desired concentration and well diluted (use vortex shakers).

8. Anesthesia. Timing: 35 min
   a) Induce deep anesthesia with an intraperitoneal (i.p.) injection of chloral hydrate (8%, 400 mg/kg; 100 µL per 10 g body weight).
   b) Leave mouse in his transport cage for 30 min.
   c) When mouse is deeply anesthetized, secure the mouse in a stereotaxic apparatus (Figure 3A).

**Alternative:** Use isoflurane. If you choose this option, ensure that the anesthesia mask is connected to an anesthesia station (MiniHUB V3, TEMSEGA, France) with two separate tubes: one for the supply gas and one for the exhaust.³⁷

Note: Supplement anesthesia as needed with a 1/3x dilution to maintain stable anesthesia throughout the experiment.

9. Craniotomy. Timing: 5 min
   a) Perform a midline scalp incision with ToughCut fine scissors
   b) Clean the area with cotton buds and sterile saline.
   c) Check the distance between bregma and lambda to ensure alignment of the skull in the anterior-posterior plane; measure left-right skull roll.
   d) Drill with a 0.9 mm steel drill bit a craniotomy above the target site(s).
Note: In our case we targeted the IPN: AP -3.3 to -3.6 mm; ML 0.35-0.5 mm; DV 4.2-4.9 mm. Use a 5º angle for all insertions into the IPN to avoid the superior sagittal sinus.

Note: The craniotomy area should be sufficiently large to allow sampling from multiple coordinates within the target brain region during acute electrophysiology recordings. However, creating an excessively large window increases the risk of brain swelling or bleeding, which can compromise both the health of the animal and the quality of the recordings. Therefore, it is important to balance accessibility with minimizing tissue damage.

10. Cannula Insertion for local infusion of MPEG₄Ch. Timing: 15 min:

a) Mount the Mini-Matrix with independently movable tetrodes and a stainless-steel injection cannula (OD 120 µm) onto the stereotaxic frame.

b) Insert the injection cannula into the brain at the target coordinates and wait 5 min for stabilization of the brain tissue.

Note: Check the mobility of tetrodes and cannula prior to surgery; replace if necessary.

Alternatives: For local infusion, you can use a glass micropipette connected to a microinjector or a One-axis Oil Hydraulic Micromanipulator.

c) To verify flow, gently expel a small drop of liquid from the cannula tip. If the cannula appears clogged, moisten the tip with filtered saline and try again. If the issue persists, replace the cannula.

d) Infuse MPEG₄Ch (250 µM; 700 nL for IPN) at a rate of 10 nL/s using the microinjector attached to the cannula.

e) Wait 5 min after infusion before retracting.

f) Verify cannula flow after the injection to confirm successful delivery.

Note: For control experiments, infuse an equivalent volume of filtered saline.

Note: For maleimide-thiol reactions performed *in vitro*, concentrations in the tens of micromolar range are typically sufficient. However, for *in vivo* applications, higher concentrations—usually in the hundreds of micromolar—are used to compensate for immediate dilution upon administration.

CRITICAL: Always confirm injection cannula placement post-hoc with histology. To check placement, apply a fluorescent dye (Dil, DiO, or DiD at the stock concentration of 1 mg/mL) to the cannula tip about 5 minutes before entering the brain (Figure 3B).

g) Wait 2 hours post-infusion before starting electrophysiological recordings.

11. Venous Catheterization (Figure 3C). Timing: 5-10 min.

Note: Venous catheterization is performed to enable precise, repeatable intravenous (i.v.) administration of nicotine (or saline) during in vivo electrophysiological recordings. This approach allows for rapid and controlled delivery of nicotine directly into the bloodstream, ensuring consistent
645 pharmacokinetics and minimizing stress or movement artifacts that could occur with repeated manual injections.

647 a) Preparation. Prepare all materials before beginning this step to minimize the duration of anesthesia and reduce risk of complications.

648 i) Fill a 1 mL syringe connected to a 30G needle with sterile saline.

649 ii) Connect the syringe to a length of PE10 tubing.

650
651
652 Note: The length of the PE10 tubing determines the volume delivered. For example, 653 16.2cm of PE10 tubing (internal radius 0.014 cm) corresponds to 10 µL, calculated as: 654
655
656
657 i) Detach the thin metal shaft of a 30G needle from its plastic base; this metal 658 shaft will be inserted into the vein.
659 ii) Insert the metal shaft of the 30G needle in the other end of the PE10 tubing.
660
661 iii) Prepare a separate 100 µL Hamilton syringe for subsequent intravenous administration of nicotine or saline.
662
663
664 b) Surgical Exposure.
665 i) Using ToughCut fine scissors, make a small incision in the skin on the inner 665 side of the hind leg to expose the underlying tissue.
666
667 ii) Carefully separate and remove the thin connective tissue layer between the 667 skin and muscle with two Dumont #5 forceps to fully expose the saphenous vein.
668
669 Note: Use fine forceps and scissors for precise dissection to avoid damaging the vein.
670
671
672 c) Catheter Insertion.
673 i) Gently insert the 30G needle shaft (connected to PE10 tubing) into the 673 saphenous vein.
674
675 ii) Slowly inject a small volume of saline to confirm that the solution flows freely 675 into the vein (no resistance or swelling at the site). Troubleshooting 4.
676
677 d) Once correct placement is confirmed, apply a small amount of SuperGlue at the entry 677 point to secure the needle and prevent movement during the experiment.
678
679
680
681
682
683
684
685
686
687
688
689
689 Caution: Avoid excess glue, which may occlude the vein or adhere to surrounding tissue.
e) Disconnect the saline syringe and connect the PE10 tubing to the prepared 100 µL Hamilton syringe containing nicotine or saline for i.v. administration during the experiment. Troubleshooting 5.
12. Electrophysiological Recordings. Timing: typically, 3–6 h
a) Before inserting the tetrodes, carefully remove the dura mater overlying the target region using a 90°-bent needle or fine forceps.

$$
Length = Volume / (\pi \times radius^2)
$$

i) Detach the thin metal shaft of a 30G needle from its plastic base; this metal shaft will be inserted into the vein.

ii) Insert the metal shaft of the 30G needle in the other end of the PE10 tubing.

iii) Prepare a separate 100 µL Hamilton syringe for subsequent intravenous administration of nicotine or saline.

b) Surgical Exposure.

i) Using ToughCut fine scissors, make a small incision in the skin on the inner side of the hind leg to expose the underlying tissue.

ii) Carefully separate and remove the thin connective tissue layer between the skin and muscle with two Dumont #5 forceps to fully expose the saphenous vein.

Note: Use fine forceps and scissors for precise dissection to avoid damaging the vein.

c) Catheter Insertion.

i) Gently insert the 30G needle shaft (connected to PE10 tubing) into the saphenous vein.

ii) Slowly inject a small volume of saline to confirm that the solution flows freely into the vein (no resistance or swelling at the site). Troubleshooting 4.

d) Once correct placement is confirmed, apply a small amount of SuperGlue at the entry point to secure the needle and prevent movement during the experiment.

Caution: Avoid excess glue, which may occlude the vein or adhere to surrounding tissue.

e) Disconnect the saline syringe and connect the PE10 tubing to the prepared 100 µL Hamilton syringe containing nicotine or saline for i.v. administration during the experiment. Troubleshooting 5.

12. Electrophysiological Recordings. Timing: typically, 3–6 h

a) Before inserting the tetrodes, carefully remove the dura mater overlying the target region using a 90°-bent needle or fine forceps.
690 **Tip:** Gently lift and cut the dura to avoid damaging underlying brain tissue. Keep the area moist with sterile saline throughout. In case of bleeding, control it with gentle pressure with hemostatic sponge.

692

693 b) **Tetrode Insertion** (Figure 3D). Timing: 30 min

694 i) Slowly lower the tetrodes into the brain at a rate of 10 µm/s until reaching the start of the region of interest (e.g., IPN).

695 ii) Once at the initial depth, wait at least 30 min to allow tissue to stabilize and reduce acute insertion effects.

696 iii) For final approach to the target region, decrease the descent speed to 1–2 µm/s.

700

701 **CRITICAL:** Slow approach and waiting period are essential to minimize tissue damage and improve recording stability.

703

704 c) Continuously monitor all channels for neuronal spikes using the acquisition software.

705 d) Adjust tetrode depth as needed to optimize signal amplitude and isolate single-unit activity. **Troubleshooting 6.**

707

708 **CRITICAL:** The most important factor is achieving a high signal-to-noise ratio and stable spike amplitudes across channels, which is crucial for reliable spike sorting.

710

711 e) **Data Acquisition:** Acquire data using a 36-channel preamplifier connected to the Neuralynx Digital Lynx 4SX data acquisition system. Record signals using Cheetah software, ensuring all channels are properly referenced and grounded.

714

715 **Alternatives:** Several alternative data acquisition systems are available for multi-channel electrophysiology. Platforms such as the open-source OpenEphys³⁸, Intan Technologies RHD recording systems, and OneBox (imec) offer cost-effective solutions that support high channel counts and are compatible with advanced probes like Neuropixels³⁹. These systems provide flexible hardware and software options for large-scale neural recordings and may be preferable for experiments requiring simultaneous monitoring of many neurons.

721

722 f) **Quality Control:** Check for stable baseline recordings and robust spike activity before proceeding with pharmacological manipulations.

723 g) If signal quality is poor (e.g., high noise, unstable amplitudes), adjust grounding, check connections, or reposition electrodes as needed.

726

727 **Note:** Always document electrode depth and coordinates for reproducibility. Maintain detailed logs of all adjustments and observations during recording.

729

730 **13. Nicotine injection:**

731 a) Begin each recording with a 5-minute baseline.

732 b) Inject either saline or nicotine i.v. in pseudo-random order. **Troubleshooting 7.**

733 c) Wait ≥5 minutes after saline and ≥15 minutes after nicotine before next injection.

734 d) For dose-response experiments, inject 7.5, 15, 30, or 60 µg/kg nicotine (2–4 injections per mouse) in pseudo-random order.

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736

737

738

739

740

14. Confirming Tetrode Placement Post-hoc:

741

742

743

744

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746

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748

749

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**CRITICAL:** Allow adequate washout between injections to avoid cumulative effects. Monitor depth of anesthesia regularly; adjust chloral hydrate accordingly.

i) To verify accurate tetrode placement after recordings, apply a small amount of a fluorescent dye (such as Dil, DiO, or DiD)—distinct from any dye used with the infusion cannula—directly onto the tetrode tip approximately 5 minutes before insertion into the brain.

ii) Following the completion of recordings, extract the brain and immerse it in 4% paraformaldehyde (PFA) at 4°C for at least one week to allow for thorough fixation.

iii) Subsequent histological processing and fluorescence imaging can then be performed to confirm the precise location of the tetrode tracks (see section 18 for detailed histology procedures).

15. Data Processing and Analysis. Timing: typically, 1–2 h per dataset, depending on channel count and software.

a) Detect extracellular spikes from raw data using CSC Spike Extractor (Neuralynx).

**Note:** Adjust detection thresholds to optimize sensitivity and minimize noise artifacts. Visual inspection of detected events is recommended.

b) Perform spike sorting using SpikeSort3D (Neuralynx), applying manual cluster cutting to isolate single units.

**CRITICAL:** Carefully inspect clusters for separation and stability. Only include well-isolated units for further analysis.

**Note:** There are many valid approaches to spike sorting; the workflow described here matches Jehl et al.,¹ but alternative open-source semi-automatic pipelines (Kilosort4⁴⁰ + Phy2, SpikeInterface,⁴¹ CellExplorer,⁴² etc.) are equally valid depending on your setup and needs. These are more efficient for high-density silicon probes (e.g., Neuropixels) or larger datasets.

c) For each well-clustered unit, extract:

i) Spike timestamps (precise timing of each detected spike)

ii) Waveform data (average or all waveforms for each unit; sometimes referred to as “waveform shapes” or “waveform matrices”)

**Note:** Consistent extraction and storage of spike times and waveforms are essential for downstream analyses.

d) Data analysis:

i) Neurons are classified as activated if their firing rate increases after nicotine administration, and as inhibited if their firing rate decreases.

ii) To determine responsiveness, a bootstrap analysis is performed by shuffling each neuron's baseline spike intervals 1,000 times to generate a distribution of expected responses.
782 iii) The actual nicotine-induced change is then compared to this distribution: a
783 neuron is considered responsive if its change in firing rate falls at or above the 98th
784 percentile (activation) or at or below the 2nd percentile (inhibition) of the shuffled data.
785 iv) For quantification, the maximal increase (for activated neurons) or minimal
786 decrease (for inhibited neurons) in firing rate is extracted within the first 3 minutes after
787 injection, and responses to nicotine are compared to those observed after saline
788 administration.
789

790 In vivo use of CATCH with behavior

791
792 This section describes how to apply the CATCH protocol to investigate the functional consequences of
793 sustained, localized receptor antagonism on animal behavior. The following procedures detail the steps
794 for animal handling and habituation, stereotaxic infusion of MPEG₄Ch, post-surgical care, and
795 behavioral testing, providing a comprehensive framework for linking molecular and cellular
796 manipulations to complex behavioral outcomes (Figure 4). This approach was used in Jehl et al.¹ to
797 reveal the role of β4 nAChR subunits in nicotine reward, but can be adapted to address a wide range of
798 neuropharmacological questions.
799
800

Note: By combining precise chemogenetic manipulation with established behavioral paradigms, such as conditioned place preference (CPP), researchers can directly assess how targeted inhibition of nAChRs in specific brain regions influences reward-related and motivational processes in vivo.

806 Timing: 5 weeks

807
808 16. **Handling and weight measurements.** Timing: 5-7 days. This procedure ensures
809 animals habituate to the experimenter and the testing environment, minimizing
810 stress-induced variability in behavioral outcomes.

811 a. Transport animals in their home cages to the behavioral testing room, or its
812 antechamber, a few days before the surgical procedure.

813 b. Scoop mouse out of cage.

814 i) Let the mouse explore your hands and arms for 8-10 minutes.

815 ii) Periodically touch the mouse head and back to habituate the mouse to
816 experimenter contact.

817 iii) Weight the mouse and return back to the cage.

818 c. Repeat procedure for all mice of the batch, changing gloves between cages/when you
819 start handling a different cage.

820 d. Repeat handling session once a day for 5 days.

821
822 Note: If needed, handling sessions can be increased to 7 days.

823
824 Note: In case the behavioral experiment includes an i.p. injection or connecting the animal to an optic
825 fiber, you can habituate it to these procedures in the two last days of handling period. For example, in
826 Jehl et al.,¹ the last 2 handling sessions comprised 8 minutes of normal handling and 2 minutes for one
827 i.p. injection of filtered saline.

828 **CRITICAL:** To reduce stress-related variability, maintain consistency between the habituation routine
829 and the actual testing procedure. During the behavioral experiment, use the same lab coat worn
830 during the handling phase. **Troubleshooting 8.**

831
832
833 **Note:** Conduct behavioral tests within the same fixed time window each day.

834
835
836
837
17. **Preparation and infusion of MPEG₄Ch for in vivo use.** Timing: 1 day. The objective is to prepare and infuse MPEG₄Ch into a target region for subsequent *in vivo* behavioral experiments.

838
a. Prior to beginning the procedure, prepare MPEG₄Ch for local injection as in step 7.

839
840 **CRITICAL:** Prepared MPEG₄Ch solution should be used within the same day. Prepare a fresh dilution
841 daily before use.

842
843
844
845
846
847
b. Pull injection glass micropipettes (Drummond 10 µL PCR Pipettes, Cat# 5-000-1001-X10) using a 2-step pull on an upright puller (Narishige PC-10) with the following parameters: Load weight: 250 g, Step 1 heat: 59.6°C, Step 2 heat: 44.9°C, Pull length: 6 mm.

848 **Note:** Store pulled capillaries in a covered plate to prevent contamination. Minimize air exposure to
849 maintain sterility. Always handle micropipettes with clean gloves.

850
851
852
853
854
855
c. Prepare for surgery by gathering all the supplies listed in the *Materials and equipment setup* section and setting up the stereotaxic apparatus (Figure 5).

853
i) Sterilize all tools in an autoclave and store them in a sterile container.

854
855
ii) Check that the isoflurane source is full and disinfect all surfaces, including the stereotaxic device.

856
iii) Ensure all reagents are available before surgery.

857
858
859
iv) Ensure that your mouse is at the appropriate age and weigh the mouse to determine the dosage of pre-anesthetic drugs.

860
861
862
863
864
865
d. Prepare the injection micropipette.

861
i) Cut the tip of the glass micropipette with sharp surgical scissors.

862
ii) Add Vaseline around the plunger.

863
iii) Fill the glass micropipette with mineral oil.

864
865
iv) Insert the plunger into the rear of the micropipette and ensure smooth ejection of mineral oil.

866
867
v) Attach the One-axis Oil Hydraulic Micromanipulator (MO-10, Narishige, Tokyo, Japan) to the stereotaxic arm and insert the micropipette.

868
vi) Ensure the plunger moves properly and fluid can be dispensed.

869
870
vii) Load the micropipette with MPEG₄Ch by touching the fluid bead on parafilm or directly from an Eppendorf tube (with risk of breaking the tip).
871 viii) To confirm accurate targeting, mix the MPEG₄Ch solution with AAV5-GFP virus (use a strong ubiquitous promoter for proper expression; for the IPN, AAV5 at a titer of 5.7 × 10¹¹ vg/mL is effective), or a fluorescent dye (Dil, DiO, DiD at the stock concentration of 1 mg/mL).

875

876 **Alternatives:** As an alternative you can use a 1µl of dye and brush the micropipette tip 5 min before entering the brain to check the micropipette track.

878

879 ix) Post hoc microscopic confirmation is required for injection site verification (see step 19).

881

882 e. Induction of anesthesia.

883 i) Place the mouse in an induction chamber and administer isoflurane at 3% with 1 L/min Oxygen, regulated with an anesthesia station (MiniHUB V3, TEMSEGA).

884 ii) Observe for deep anesthesia before transferring the mouse to the surgical platform.

887 iii) Shave the top of the head in two passes: first in the direction of hair growth, then against it.

888 iv) Remove all loose hair.

890 v) Secure the mouse in the stereotaxic frame by gently pulling the tongue forward.

891 vi) Position the incisor bar and connect the respiratory tube.

892 **Alternatives:** Other anesthetics, such as ketamine/xylazine mix, can be used in place of isoflurane.

893

894 f. Maintain anesthesia with isoflurane at 1.5% with 0.5 L/min Oxygen.

895

896 **Note:** If the mouse starts gasping, reduce isoflurane concentration.

897

898 g. Maintain body temperature with a heating mat at 37°C.

899 h. Surgical Preparation.

900 i) Inject Lidocaine (100 µL) under the skull's skin at the incision site.

901 ii) Administer Buprenorphine (1 mg/kg) 30 min before the end of surgery.

902 iii) Apply physiological gel to the eyes to prevent dryness.

903 i. Mark the mouse for identification: Left ear → Mouse 1, Right ear → Mouse 2, Both ears → Mouse 3, etc.

904 j. Disinfect the incision site with betadine and dry with a sterile cotton bud.

906 k. Stereotaxic surgical preparation.

907 i) Make a midline incision from the eyes to the skull base with a fine scissor.

908 ii) Remove conjunctival tissue from the skull surface.

909 iii) Identify bregma and lambda as stereotaxic reference points.

910 iv) Ensure the skull is level with bregma and lambda before proceeding. If these are <0.05 mm different, you can proceed. If it is >0.05 mm different, adjust the mouse's placement in the ear bars or the height of the bite bar until the skull is flat in the AP axis.

914

915 **Note:** AP distance between bregma and lambda should be around 4.2mm in an adult C57 mouse.
916 I. Mark the target site using stereotaxic coordinates.

917

918 **Note:** For the IPN we used the following target: AP: +3.4 mm, ML: ±0.4 mm. Angle: 5°.

919

920 m. Drill through the skull at the marked site using a 0.9 mm steel drill bit, avoiding dura damage.

922 n. Injection.

923 i) To verify flow, gently expel a small drop of liquid from the micropipette tip. If the pipette appears clogged, moisten the tip with filtered saline and try again.

925

926 **Alternative:** Alternatively, carefully shorten or break the tip using forceps to restore flow. If the issue persists, replace the micropipette.

928

929 ii) Lower the micropipette to the target depth (DV axis).

930 iii) Wait 2 min before injecting 500 nL at 100 nL/min (5 full turns on the One-axis Oil Hydraulic micromanipulator).

932 iv) After injection wait 4 min, withdraw the needle by 0.1 mm, and wait another 1 min before slowly withdrawing the needle while monitoring for fluid leakage. (Total waiting time = 5 min).

935 v) Verify pipette flow after the injection to confirm successful delivery.

936 o. Fixation and Closure.

937 i) Rehydrate the skull with saline (do not dry or remove excess).

938 ii) If the procedure is lengthy, inject 0.9% saline subcutaneously.

939 iii) Suture the scalp using 3-4 stitches.

940 iv) Clean the incision area with betadine.

941 p. Turn off the isoflurane and remove ear bars carefully.

942 q. Post-Surgical Care.

943 i) Place the mouse on a heating pad for recovery.

944 ii) Monitor the mouse until fully awake.

945 iii) Check for signs of distress or complications.

946 iv) Ensure proper hydration and administer analgesics as per protocol.

947 v) Verify carbon filter weight and isoflurane levels before ending the session.

948 vi) Monitor mouse during the three days following surgery.

949

950 18. Behavioral task: Conditioned Place Preference (CPP) test. Timing: 5 days.

951

952 **Note:** The following step describes the conditioned place preference (CPP) task that we performed in Jehl et al.¹ We used this to investigate the role of β4 nAChR subunits in the regulation of nicotine reward. However, the protocol can be adjusted for other behavioral tests.

955

956 **CRITICAL:** In Jehl et al.,¹ the CPP test in KI mice was initiated three days after MPEG₄Ch infusion into the IPN. We recommend allowing a three-day interval between infusion and behavioral testing.

958 However, ex vivo data from the IPN indicate that MPEG₄Ch-mediated antagonism persists for at least 10 days post-infusion. This suggests that a longer interval may be possible but should be empirically validated.
961 **Note:** To minimize stress, avoid changing the cages of experimental animals during the same week as
962 behavioral testing.

963 a. Prepare the behavioral testing environment by arranging all necessary equipment
964 including:

965 i) Dimmable panel light.

966 ii) CPP chamber.

967 **Note:** We used a Y-maze apparatus (Imetronic, Pessac, France) with one arm closed and the two
968 remaining arms fitted with manually operated doors. This setup created two rectangular chambers
969 (11 x 25 cm), each featuring distinct cues in texture and color, separated by a central triangular
970 neutral compartment (11 cm per side). One chamber had grey textured floors and walls, while the
971 other had smooth floors and walls with black and white stripes.

972 **CRITICAL:** The combination of textures and colors chosen for each chamber were balanced in previous
973 pilot experiments to minimize mice's initial preferences. [Troubleshooting 9](#).

974 iii) An overhead video camera (Intel RealSense, 640x480, 30 frames per s).

975 iv) A computer with recording software for video acquisition.

976 **CRITICAL:** Light intensity during testing significantly influences behavioral outcomes. In this protocol,
977 light levels were maintained at 100 lux within the CPP chambers.

978 **CRITICAL:** Animals should be transferred to the testing room or holding area at least 30 minutes prior
979 to each behavioral session (from Day 1 to Day 5) to allow acclimatization.

980 b. Pretest, day 1:

981 i) Take each mouse individually and place it in the middle chamber of the CPP
982 apparatus.

983 **Note:** In this pretest session, allow the mouse to freely explore and access both open arms of the Y
984 maze
985 ii) Record with the overhead video camera the activity of the animal for 15 minutes.

986 iii) After placing the mouse back in its cage, thoroughly clean the apparatus using
987 Surfa'Safe Premium disinfectant spray (Laboratoires Anios, #20003N). Change
988 gloves before handling the next mouse, and repeat this process for all animals.

989 iv) After the pretest session, select a video analysis method to detect the animal's
990 position and calculate its trajectories and time spent in each chamber.

992 **Note:** In Jehl et al.,¹ we used a custom-made Python recording software to record the video, and
993 ezTrack³³ to track the position of the animal offline. Alternatively, open-source Bonsai⁴³,⁴⁴, or
994 EthoVision and ANY-maze software can be used for behavioral tracking.

996
997 v) Once the time spent in the two chambers is determined, randomly assign
998 individual mice into two balanced groups. In one group, nicotine is paired with
one chamber, and in the other group, nicotine is paired with the opposite chamber to ensure an unbiased CPP design.

c. Conditioning days, day 2 to 4:

i) Prepare filtered saline (0.9% NaCl) and nicotine tartrate solution (0.2 mg/kg, in filtered saline) as described in *Materials and equipment setup* section.

**Note:** Each conditioning day includes two conditioning sessions, at least 5 hours apart from each other. In one conditioning session an i.p. injection of nicotine is paired with one chamber, whereas in the other session a saline injection is paired with the other chamber. The order of saline - nicotine session is counterbalanced across the 3 conditioning days.

**CRITICAL:** A saline-saline control group, in which both chambers are paired with saline should be added as a control.

ii) Scoop mouse out of its home-cage and perform an i.p. injection of either saline or nicotine solution.

iii) Immediately after injection, place the animal into the appropriate chamber for 20 minutes (videorecording these sessions is not necessary).

iv) After placing the mouse back in its cage, thoroughly clean the apparatus with a disinfectant spray. Change gloves before handling the next animal, and repeat this process for all mice.

v) Repeat [c.ii, c.iii, c.iv] for all conditioning sessions on all conditioning days.

d. Test day, day 5: repeat procedure of pretest [b.i-iv].

e. Behavioral analysis: The preference score (in seconds) is calculated by subtracting the pretest difference from the test difference using the following formula:

$$ (tNIC - tSAL)TEST - (tNIC - tSAL)PRE $$

where:

tNIC = time spent in the nicotine-paired chamber

tSAL = time spent in the saline-paired chamber

TEST = values recorded on the test day

PRE = values recorded on the pretest day

This formula quantifies the change in preference for the nicotine-paired chamber

from pretest to test. **Troubleshooting 10 and 11**

19. Post-hoc verification of *in vivo* injection and recording sites

a) Perform intracardiac perfusion with 1X PBS followed with 4% PFA to fix the brain (must be done inside a chemical fume hood).

b) Post-fix the brain in 4% PFA at 4°C for at least 3 days.

c) Section the fixed brain coronally into 50 µm thick slices using a vibratome (Leica VT1000S).

999

1000

1001

1002

1003

1004

1005

1006

1007

1008

1009

1010

1011

1012

1013

1014

1015

1016

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1021

1022

1023

1024

1025

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1028

1029

1030

1031

1032

1033

1034

1035

1036

1037

1038

1039

1040

1041

1042
1043 d) For verification of injection or tetrode placement, directly visualize fluorescent dye (e.g., Dil, DiO, DiD) in brain sections using epifluorescence microscopy.

1044 [Troubleshooting 12.](#)

1045 e) If AAV5-GFP virus or other immunolabeling is used, perform immunohistochemistry with an anti-GFP antibody as needed.

1046 f) Image sections at 10x magnification with an epifluorescence microscope to confirm the location of MPEG₄Ch infusion or recording sites.

1047 g) To visualize the slice images, use the FIJI³⁴ software. Save the images as TIFF files for long-term storage.

1048 h) For whole-brain anatomical mapping, use DMC-BrainMap³⁵, an open-source napari plugin, to register injection sites to the Allen Brain Atlas.

1049

## 1050 Expected outcomes

1051 When performed as described, this protocol enables robust and reproducible assessment of CATCH-mediated receptor antagonism at multiple levels. *Ex vivo*, acute brain slice recordings from Chrnb4E61C-KI mice treated with MPEG₄Ch (50 µM, 10 min) show a pronounced and sustained reduction in nicotine-evoked whole-cell currents. In Jehl et al.,¹ MPEG₄Ch-treated slices exhibited an average reduction of over 80% in peak nicotine-induced current amplitude compared to untreated controls, confirming the efficacy and specificity of the approach (Figure 6A). This antagonism was selective for the targeted nAChR subunit and persisted for the duration of the recording, while untreated slices maintained stable, robust responses.

1052 In vivo, local infusion of MPEG₄Ch results in sustained antagonism of nAChRs, as evidenced by altered neuronal firing patterns in electrophysiological recordings. Neurons in the target region display either reduced or abolished nicotine-induced activation, and these changes can be quantitatively analyzed using the described spike sorting and statistical methods (Figure 6B). At the behavioral level, animals subjected to CATCH manipulation exhibit altered responses in paradigms such as conditioned place preference (CPP), reflecting the functional consequences of localized, sustained receptor blockade (Figure 6C).

1053 Together, these outcomes provide a comprehensive demonstration of the specificity, efficacy, and functional impact of the CATCH approach, serving as a benchmark for users to evaluate the success of their own experiments.

1054

1055

## 1056 Limitations

1057 While this protocol offers powerful and precise control over receptor function, several limitations should be considered.

1058 This protocol is optimized for use with the Chrnb4E61C KI mouse line, which ensures selective and stable expression of the engineered receptor subunit throughout development and across relevant brain regions. While this genetic approach provides high specificity and reproducibility, it does limit the
1086 method's immediate applicability to other receptor types or species unless analogous KI lines are
1087 available. Generating new KI lines can be time-consuming and resource-intensive.

1088 As an alternative, viral vectors can be used to express cysteine-mutated receptors in specific brain
1089 regions or cell types; however, this approach may result in incomplete replacement of native subunits
1090 and variable expression levels, potentially reducing the precision and consistency of receptor targeting.

1092
1093 **Troubleshooting**

1094
1095 **Problem 1:**

1096 Unhealthy brain slices.
1097 In whole-cell patch-clamp recordings (step 3), slice health is critical for stable and reliable data.
1098 Unhealthy or deteriorating slices may result from poor slicing conditions, excessive delay after dissection, or inadequate maintenance of physiological conditions during the experiment. This can lead to unstable recordings or failure to obtain usable data.

1101
1102 **Potential solution:**

1103
1104 Perform transcardiac perfusion efficiently to minimize ischemia, and keep the cutting solution ice-cold and well-oxygenated throughout dissection and slicing.

1105 Record only from slices within the first 6 hours after dissection, when tissue viability is highest.

1107
1108 Continuously bubble all solutions, including MPEG₄Ch-containing 1X aCSF, with carbogen gas to prevent hypoxia.

1109
1110 If issues persist, thoroughly clean all experimental materials (slicing tools, chambers, tubing, reservoirs, perfusion lines) and re-prepare all solutions to eliminate possible contamination or degradation.

1112
1113 **Problem 2:**

1114 No response to nicotine puffs (step 4).
1115 Possible causes include a clogged puff pipette, low nicotine concentration, or failure to deliver the solution.

1117
1118 **Potential solution:**

1119 Test puff delivery outside the slice before recording. If blocked, refill and clean the lines with filtered 1x aCSF or ethanol followed by Milli-Q water. Check the pneumatic pump pressure. If the problem persists, prepare a fresh nicotine solution.

1122
1123 **Problem 3:**

1124 No change in nicotine-evoked current amplitude after MPEG₄Ch incubation (step 5).
1125 Possible causes include lack of receptor expression in the recorded neurons, degraded MPEG₄Ch aliquots, incorrect stock concentration or dilution errors.

1127
<figure><img src="image_5.png" /></figure>

1128 Potential solution:

1129 Verify receptor expression in the target areas or cells using available assays, such as
1130 immunostaining.

1131 Include positive control experiments in brain regions with confirmed receptor expression.

1132 Assess MPEG₄Ch integrity using high-performance liquid chromatography (HPLC), optionally
1133 coupled with mass spectrometry.

1134 Use freshly prepared aliquots and dilute only into freshly oxygenated 1x aCSF immediately
1135 before use. Re-prepare the working or stock solution if needed.

1136

1137 Problem 4:

1138 Difficulty with saphenous vein catheterization (vein collapse, poor visualization, bleeding, swelling, or unsuccessful insertion; step 11).

1139 Possible causes include dehydration, vasoconstriction, insufficient exposure, accidental puncture through both sides of the vein, rough handling, or the needle not being in the vein lumen.

1142

1143 Potential solution:

1144 Warm the limb to dilate the vein and ensure the animal is well-hydrated.

1145 Carefully dissect and fully expose the vein; use a stereomicroscope if needed.

1146 Use gentle, shallow insertion to avoid puncturing through the vein.

1147 If bleeding or swelling occurs, apply gentle pressure with a cotton swab and try a new site along the vein or switch to the other limb.

1148 If saline does not flow freely, withdraw slightly and reinsert the needle, check for kinks or blockages in the tubing, and flush tubing before insertion.

1151

1152 Problem 5:

1153 Catheter instability, leakage, or loss of patency during the experiment (step 11).

1154 Possible causes include inadequate fixation, animal movement, glue not set, poor seal, or excessive movement/tension on the tubing.

1156

1157 Potential solution:

1158 Hold the needle steady until the glue sets and use only a minimal amount to avoid occlusion.

1159 Ensure the animal is immobilized and the tubing is not under tension.

1160 Reapply glue if leakage occurs and check fixation before proceeding.

1161 Minimize movement during the experiment to maintain catheter patency.

1162

1163 Problem 6:

1164 No or weak signal high noise, or unstable baseline during *in vivo* electrophysiology (step 12).

1165 Possible causes include poor electrode placement, blocked or damaged electrodes, tissue damage, poor grounding, electrical interference, anesthesia fluctuations, animal movement, electrode drift, or tissue swelling.
1168

### Potential solution:

1170 Verify stereotaxic coordinates and carefully check tetrode integrity before insertion;
1171 reposition the probe if needed and allow tissue to settle after insertion.

1172 Ensure all ground and reference connections are secure; minimize cable and animal
1173 movement; shield the setup from external electrical noise.

1174 Monitor and maintain stable anesthesia throughout the recording.

1175

### Problem 7:

1177 Nicotine response is absent or blunted in both control and MPEG<sub>4</sub>Ch conditions (step 13).

1178 Possible causes include incorrect catheter placement, degraded nicotine solution, or anesthesia depth
1179 suppressing neuronal responsiveness.

1180

### Potential solution:

1182 Verify intravenous catheter patency before each injection.

1183 Prepare fresh nicotine solution and confirm correct dosing.

1184 Monitor anesthesia depth and adjust as needed to maintain physiological responsiveness.

1185

### Problem 8:

1187 Mice remain anxious or show high stress during handling or testing (step 16).

1188 Possible causes include inconsistent handling routine, insufficient habituation, handling by multiple
1189 experimenters, or sudden environmental changes.

1190

### Potential solution:

1192 Ensure handling is performed daily at the same time, by the same experimenter, and in a calm
1193 environment. Increase the number of habituation days if needed. Minimize noise and disturbances.

1194 Use the same lab coat and gloves as during testing.

1195

### Problem 9:

1197 All mice prefer one chamber in the pretest (CPP; step 18).

1198 Possible causes include unbalanced cues (texture, color, light) or strong innate preference.

1199

### Potential solution:

1201 Pilot test chamber cues and balance them across groups. Adjust features such as color and texture
1202 combinations to minimize innate preference. Randomize the assignment of drug-paired chambers for

1203 each animal.

1204

### Problem 10:

1206 No behavioral effect of CATCH manipulation (step 18e).

1207 Possible causes include ineffective infusion (missed target, degraded compound), insufficient receptor
1208 blockade, or absence of the target receptor.
1209

1210

### Potential solution:

1211

* Confirm infusion site with histology.

1212

* Use freshly prepared MPEG₄Ch solution.

1213

* Confirm receptor targeting and blockade with ex vivo or in vivo electrophysiology in a subset of animals.

1214

1215

* Consider increasing the MPEG₄Ch concentration or adjusting the timing between infusion and behavioral testing.

1216

1217

* Include positive control experiments targeting brain regions with confirmed receptor expression.

1218

1219

1220

### Problem 11:

1221

Off-target effects or unexpected behavioral/physiological changes after MPEG₄Ch infusion (step 18e).

1222

Possible causes include diffusion of MPEG₄Ch outside the intended region, excessive volume, or non-specific binding.

1223

1224

1225

### Potential solution:

1226

* Reduce infusion volume or slow the infusion rate to minimize spread.

1227

* Use a fluorescent dye to visualize the extent of spread post-hoc and adjust parameters accordingly.

1228

1229

* Confirm that the observed effects are absent in wild-type or non-targeted animals to rule out non-specific actions.

1230

1231

* Consider using a control ligand or vehicle infusion as an additional control.

1232

1233

### Problem 12:

1234

No dye or marker detected at the intended site during histology (step 19).

1235

Possible causes include dye washed out during processing, incorrect dye used, incorrect loading of mix of MPEG₄Ch, clogging of the pipette, or insufficient fixation.

1236

1237

1238

### Potential solution:

1239

Use appropriate dye concentration and fixation protocol. Check carefully when loading solution in pipette and check flow of solution in pipette right before lowering it to inject. Solution should be vortexed right before loading of the pipette to preserve its homogeneity. Minimize time between brain extraction and fixation. Use a dye distinct from other markers. Protect dye from light.

1240

1241

1242

1243

1244

### Resource availability

1245

**Lead contact:** Further information should be directed to and will be fulfilled by the lead contact, Nicolas Guyon (nicolas.guyon@espci.fr).

1246

1247

**Technical contact:** Technical questions on executing this protocol and requests for resources and reagents should be directed to and will be answered by the technical contact, Alexandre Mourot (alexandre.mourot@espci.fr).

1248

1249
1250 **Materials availability:** The KI mouse line Chrnb4<sup>E61C</sup> and the covalent antagonist MPEG<sub>4</sub>Ch generated
1251 in this study will be made available on request, but we may require a completed materials transfer
1252 agreement. Additionally, MPEG<sub>4</sub>Ch is commercially available at Broadpharm® (San Diego, CA, USA).

1253 **Data and code availability:** This study did not report datasets or original code.

1254

1255 Acknowledgments

1256 The authors would like to thank Joachim Jehl, Eléonore Vicq, Hélène Geoffroy, as well as Otilia de
1257 Oliveira and Emilie Tubeuf at the animal facility at ESPCI Paris, for their assistance. We thank DIM C-
1258 BRAINS (Région Ile-de-France) and NeuroPSL (Université PSL) for their support. Image of beakers<sup>45</sup>
1259 and mouse<sup>46</sup> in graphical abstract and of brain<sup>47</sup> in figure 4 were adapted from SciDraw
1260 (https://scidraw.io/).

1261
1262
1263 This work was funded by the Agence Nationale de la Recherche (ANR-21-CE16-0012 CHOLHAB to A.M. and ANR-21-CE37-0026 NICOPTOTOUCH to A.M. and ANR-24-CE16-6957 NIMONAR to A.M.); the Human Frontier Science Program (RGP0035/2020 to A.M.); the National Institutes of Health (NS111600 to A.M.); the Fondation pour la Recherche Médicale (Grant EQU201903007961 to A.M. and a fourth-year PhD fellowship FDT202404018105 to M.C.); and by a Swedish Research Council Post-doctoral fellowship (2022-06168 to N.G.).
1264
1265
1266
1267
1268

1269 Author contributions

1270 A.M. and N.G. conceptualized and supervised the protocol. A.M. provided the resources. M.C., K.S.,
1271 A.M., and N.G. contributed to writing the original draft, and were involved in writing, reviewing, and
1272 editing the manuscript. M.C., K.S., A.M., and N.G. contributed to the visualization. A.M. and N.G.
1273
1274 acquired the funding.

1275 Declaration of interests

1276 The authors declare no competing interests.

1277

1278 Declaration of generative AI and AI-assisted technologies in the
1279 writing process

1280 During the preparation of this work the authors used Sana AI (Sana Labs) and ChatGPT in order to
1281 improve language and readability, with caution. After using this tool, the authors reviewed and edited
1282 the content as needed and take full responsibility for the content of the published article.
1283
1284

1285

1285 References

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1435
1436 Figure legends
1437
1438 Figure 1: The CATCH method.
1439 A) In CATCH, a genetically-modified receptor incorporates a cysteine residue, for the covalent anchor-
ing of a cysteine reactive antagonist. The tethered ligand is composed of a cysteine-reactive group, a
1440 flexible linker and a functional antagonist. After conjugation, the modified receptor becomes irreversi-
1441 bly antagonized.
1442
1443 B) Chemical structure of MPEG₄Ch.
1444 C) Chemical structures of MPEG derivatives with various PEG lengths (n) and headgroups, available
1445 from Broadpharm®. Alkyne groups can be used for attaching more complex ligands through high-yield
1446 click chemistry.⁴⁸ Example structures and additional information are available at:
1447 https://broadpharm.com/product-categories/Thiol-Reactive-Linkers/maleimide-linkers#mal-amido-
1448 peg-alkyne.

1449 D) High resolution structure of the human α3β4 nAChR in complex with nicotine (PDB code 6pv7).
1450 Only a dimer is shown for clarity. Inset, zoom in on the ACh binding pocket, showing docked nicotine
1451 and amino acid E61 at close proximity, the position mutated to cysteine.

1452 E) β4E61C-containing nAChRs are irreversibly antagonized after conjugation with MPEG4Ch, but not
1453 wild-type (WT) receptors.

1454
1455 Figure 2: *Ex vivo* electrophysiological recordings with CATCH.

1456 A) Acute brain slices are incubated with 50 µM of MPEG₄Ch for 10 minutes under oxygenation, then
1457 rinsed three times with fresh 1X aCSF before immediate transfer to the patch clamp chamber.

1458 B) Alternatively, MPEG₄Ch can be injected *in vivo* in KI mice, allowing direct patch-clamp recordings
1459 from acute slices without further incubation.

1460
1461 Figure 3: *In vivo* acute electrophysiological recordings with CATCH.

1462 A) Stereotaxic frame equipped with a Thomas Recording (TRec) Matrix, enabling precise injection and recording with multiple independently movable tetrodes.

1463 B) Application of fluorescent dyes to the tip of a glass micropipette (or injection cannula) using a P2 pipette. This allows for post-hoc verification of the glass micropipette track to confirm accurate targeting of the brain region.

1467 C) Catheterization of the saphenous vein for i.v. drug administration, such as nicotine. A needle tip attached to a plastic tube is inserted into the vein close to an 90° angle to avoid perforation. Adhesive tape on the paw and pipette tips placed behind the hind limb help to tense the leg, facilitating easier access to the saphenous vein.

1471 D) Lowering the tetrodes into the brain following craniotomy and removal of the dura mater.

1472
1473 Figure 4: Experimental workflow for the *in vivo* use of CATCH with behavior.
1474 Figure 5: Stereotaxic surgery setup.
1475 A) An upright micropipette puller, used with thin-walled glass capillaries to produce glass pipettes
1476 suitable for intracranial injections.
1477 B) Example of a typical stereotaxic setup, including surgical tools, a stereotaxic frame with ear bars
1478 and a nose holder for head immobilization and gas anesthesia, and a control panel for coordinate recording.
1479 The oil hydraulic micromanipulator is attached to a manipulator arm to ensure precise intra-
1480 cranial injections. A heat pad maintains body temperature under anesthesia, while a stereomicro-
1481 scope provides magnified visualization of the surgical field.
1482
1483 Figure 6: Expected results.
1484 A) *In ex vivo* slices, local infusion of MPEG₄Ch in Chrnb4E61C mice inhibited nicotinic current in IPN neurons.
1485
1486 B) *In vivo* electrophysiological recordings show reduced nicotine-induced activation of IPN neurons
1487 after MPEG₄Ch infusion in Chrnb4E61C mice. Representative immunofluorescence image showing an
1488 injection track labeled with fluorescent dye (left), and GFP expression confirming correct targeting in
1489 the IPN (right). Scale bars: 500 µm.
1490
1491 C) In a conditioned place preference (CPP) test, blockade of β4 nAChR subunits in the IPN via
1492 MPEG₄Ch infusion increases preference for a low nicotine dose (0.2 mg/kg i.p) in Chrnb4E61C mice.
1493 Significant (*) is defined here as p < 0.05. Figure reprinted and adapted with permission from Jehl et
1494 al., 2025¹.
<figure><img src="image_9.png" /><figcaption>Figure 1</figcaption></figure>

<figure><img src="image_10.png" /></figure>

<figure><img src="image_11.png" /></figure>

<figure><img src="image_12.png" /></figure>

<figure><img src="image_13.png" /></figure>
<caption>Figure 2</caption>

Incubation with
MPEG₄Ch for 10 min

<figure><img src="image_14.png" /></figure>

Patch clamp recordings
in the IPN

<figure><img src="image_15.png" /></figure>

<figure><img src="image_16.png" /></figure>

B Local MPEG₄Ch injection in the IPN

<figure><img src="image_17.png" /></figure>

<figure><img src="image_18.png" /></figure>
A

Stereotaxic
control panel

<figure><img src="image_19.png" /></figure>

B

P2 pipette +
filter tips

<figure><img src="image_20.png" /></figure>

<figure><img src="image_21.png" /></figure>

Fluorescent dyes
(Dil, DiO, DiD)

<figure><img src="image_22.png" /></figure>
<figure><img src="image_23.png" /><figcaption>Figure 4</figcaption></figure>
<figure><img src="image_24.png" /><figcaption>Figure 5</figcaption></figure>
<figure><img src="image_25.png" /><figcaption>A. Patch clamp recordings</figcaption></figure>

β4<sup>E61C</sup>
mouse

Pre-incubation
with MPEG₄Ch

<figure><img src="image_26.png" /></figure>

<figure><img src="image_27.png" /></figure>

B

Tetrode
recording

<figure><img src="image_28.png" /></figure>

β4<sup>E61C</sup>
mouse

MPEG₄Ch
infusion + Dye
or AAV-GFP

<figure><img src="image_29.png" /></figure>

<figure><img src="image_30.png" /></figure>

<figure><img src="image_31.png" /></figure>

C

β4<sup>E61C</sup>
mouse

<figure><img src="image_32.png" /></figure>

MPEG₄Ch
infusion

<figure><img src="image_33.png" /></figure>

<figure><img src="image_34.png" /></figure>