# Hijacking a real time detection thermocycler for enzymology: Improvement of a fluorescent bulk assay monitoring helicase activity

Jean-Philippe Robin, Vincent Mocquet

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Jean-Philippe Robin, Vincent Mocquet. Hijacking a real time detection thermocycler for enzymology: Improvement of a fluorescent bulk assay monitoring helicase activity. Biochimie, In press, 10.1016/j.biochi.2025.02.002. hal-05069612

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# Hijacking a real time detection thermocycler for enzymology: Improvement of a fluorescent bulk assay monitoring helicase activity

Jean-Philippe Robin, Vincent Mocquet *

Laboratoire de Biologie et Modelisation de la Cellule, Ecole Normale Superieure de Lyon, CNRS, UMR 5239, Inserm, U1293, Universite Claude Bernard Lyon 1, 46 allee d'Italie F-69364 Lyon, France

## ARTICLE INFO

**Article history:**
Received 19 July 2024
Received in revised form
10 December 2024
Accepted 17 February 2025
Available online xxx

Handling Editor: B Friguet

**Keywords:**
Helicase assay
Fluorescent probe
UPF1
Real time detection thermocycler

## ABSTRACT

Helicases are enzymes involved in all aspects of nucleic acid synthesis, regulation and degradation. As a consequence, several methods were developed to monitor their enzymatic activity. In this report, we described an improvement of bulk fluorescent helicase assays to overcome their specific limitations (cost, health and safety regulations, etc.). Using a real time detection thermocycler to monitor the fluorescence in real-time, we managed to precisely control the initiation of the helicase reaction through temperature tuning. Therefore, we were able to demonstrate that this setup could provide a qualitative and a quantitative evaluation of the helicase domain of the UPF1 helicase (UPF1-HD) and that several fluorophores could be used in parallel during the same run.

© 2025 The Authors. Published by Elsevier B.V. This is an open access article under the CC BY license (http://creativecommons.org/licenses/by/4.0/).

## 1. Introduction

Helicases are molecular motors that use the energy derived from binding and hydrolysis of nucleoside triphosphate (typically ATP) to translocate on DNA or RNA and disrupt hydrogen bonds between complementary strands of the DNA or RNA double helix. Some helicases (e.g., DEAD box RNA helicases) only unwind local patches of nucleic acids. Others (e.g., UPF1-like) are processive: they translocate onto longer segments of single stranded nucleic acids, unwind double stranded structures and displace nucleic-acid-bound proteins. This translocation can be 5'-to-3' or 3'-to-5' orientated [1-3]. Helicases play crucial roles in all aspects of nucleic acid metabolism, from DNA replication to RNA decay, including transcription, DNA repair, telomere maintenance, splicing, export and translation. Highlighting their overall importance, the inhibition of their activities due to point mutations or exogenous stress is linked to cancer development, neurological pathologies, autoimmune diseases or developmental defects [4-8]. Therefore, helicases now represent a significant drug target class and identifying

small molecules with specific helicase inhibitory effects for cancer or antiviral therapy is the subject of intense research [9-13].

Various techniques have been developed to monitor helicases activities, broadly categorized into bulk assays and single-molecule assays, each with distinct advantages and limitations [14]. Single-molecule assays provide high resolution data. For example, magnetic tweezer and optical tweezer measure the mechanical forces exerted by a single molecule of a helicase on a single molecule of its nucleic acid substrate, providing mechanistic insights into the helicase activity [15,16]. Similarly, single-molecule Förster Resonance Energy Transfer (FRET) assays utilize the energy-transfer interaction between a pair of reporter molecules and measure distance changes on a range of a few nanometers, allowing probing different functional aspects of helicases [17,18]. In addition, combining both approaches (mechanical force measuring and FRET) is also feasible [19]. However, these techniques use complicated experimental setups unavailable to many laboratories and require specialized expertise to interpret the assays. Alternatively, bulk assays are simpler, widely accessible and less costly. As an output, they provide measurements averaged from a population of helicase molecules and therefore are not well suited to interrogate conformation dynamics. Nevertheless, these approaches are perfectly compatible with "proof-of-concept" studies on probes conformation, chemical inhibitors or helicase mutations.

<footnote>This article is part of a special issue entitled: Structure Integration, Function, and Reactivity of RNA published in Biochimie.</footnote>

<footnote>* Corresponding author.
E-mail address: vincent.mocquet@ens-lyon.fr (V. Mocquet).</footnote>
Traditional bulk helicase assays often employ radiolabelled probes, where the helicase-mediated unwound probe can be detected via autoradiography or scintillation counting [20–22]. These assays offer high sensitivity but come with significant drawbacks: (1) they typically provide only end-point data, lacking real-time dynamics of helicase activity; (2) handling and disposal of radioactive materials pose safety risks and environmental concerns; (3) strict regulations for the use of radioactive substances may increase operational complexity and cost. By contrast, bulk fluorescence-based assays provide a way to measure the overall helicase activity by measuring changes in fluorescence, which correlate with the unwinding of the nucleic acid substrate (either with fluorescent intercalating dyes or with the combination of a fluorescent probes and a quencher molecule relying on FRET) [23–27]. These approaches have the advantage of not using hazardous radioisotope molecules and the possibility to monitor real time activity when combined with the adequate equipment. Unfortunately, spectrofluorometers typically used for these assays are prohibitively expensive for many laboratories, while many affordable systems lack a precise temperature control, which it is yet crucial for maintaining enzyme activity and assay consistency, especially for real-time monitoring.

Here, we improve the existing bulk fluorescent helicase assay by employing a real-time **detection thermocycler** system and triggering the helicase activity by temperature control. Our setup enables sensitive and real-time acquisition of fluorescence with different wavelengths and with a precise temperature control. Moreover, our assay is performed in a 96-well plate format, thereby reducing reaction volumes compared to standard fluorescence protocols and importantly facilitating high-throughput analysis. In addition, we developed a custom-made R script to interpret the generated data. We benchmark our approach by measuring the helicase activity of Up-frameshift 1 (UPF1). UPF1 is a multifunctional enzyme involved in different DNA- and RNA-related processes occurring in the nucleus as well as in the cytoplasm [28]. The best documented function of UPF1 is its role in the Nonsense-mediated MRNA Decay (NMD). Following ATP hydrolysis, the two RecA domains that constitute UPF1 helicase domain (HD) undergo a conformational change, allowing the translocation of the enzyme from the 5' to the 3' end of the substrate only. Thereby, UPF1-HD can unwind the double stranded DNA or RNA substrates structures that will be faced during translocation [15]. Thus, in this article, we prove that our strategy is adapted to efficiently measure UPF1 helicase activity and quantitatively discriminate the effects of UPF1 mutations, exogenous inhibitors and DNA probes structures on its activity.

## 2. Material and methods

### 2.1. Protein purification

UPF1 helicase domain (UPF1-HD) WT or mutant forms, as well as the UPF1-HD/Tax or UPF1-HD/GST complexes were produced and purified exactly as previously described [29].

### 2.2. UV crosslinking

The reaction mix containing proteins and DNA oligos was prepared according to the “DNA probe formation” and “unwinding assay” procedure described below. Subsequently, the reaction mix was placed into a UV crosslinker (Stratagene Stratalinker UV 1800 Crosslinker) and exposed to UV light at a dose of 500 mJ/cm². Finally, sample migration was performed on a 5 % polyacrylamide gel to evaluate the protein/probe binding.

### 2.3. DNA probes

The different DNA duplexes were formed by the hybridisation of the following oligos (Integrated DNA Technologies, IDT™), as described in the figures.

Setup-I.

- ps32: TCA CTA TAG GGA CAC AAA ACA AAA GAC AAA AAG TGT GCG TAC AAC TAG CT

- fluo1:/5FAM_6/AGC TAG TTG TAC GCA CAC

complementary oligo: GTG TGC GTA CAA CTA GCT/3BHQ_1/ Setup-II (construction (i))

- ps32: TCA CTA TAG GGA CAC AAA ACA AAA GAC AAA AAG TGT GCG TAC AAC TAG CT/3BHQ_1/

- fluo1:/5FAM_6/AGC TAG TTG TAC GCA CAC

complementary oligo: GTG TGC GTA CAA CTA GCT.

**Construction (ii).**

- ps32: TCA CTA TAG GGA CAC AAA ACA AAA GAC AAA AAG TGT GCG TAC AAC TAG CT/3BHQ_1/

- fluo1:/5FAM_6/AGC TAG TTG TAC GCA CAC

- 18mer middle: TTT TTG TCT TTT GTT TTG

complementary oligo: GTG TGC GTA CAA CTA GCT.

**Construction (iii) and RNA probe.**

- ps14: TCA CTA TAG GGA CAG TGT GCG TAC AAC TAG CT/3BHQ_1/

- RNA probe: UCA CUA UAG GGA CAG UGU GCG UAC AAC UAG CU/ 3BHQ_1/

- fluo1:/5FAM_6/AGC TAG TTG TAC GCA CAC

complementary oligo: GTG TGC GTA CAA CTA GCT.

**Construction (iv).**

- ps32: TCA CTA TAG GGA CAC AAA ACA AAA GAC AAA AAG TGT GCG TAC AAC TAG CT/3BHQ_1/

- fluo1:/5FAM_6/AGC TAG TTG TAC GCA CAC

- 14mer beginning: TGT CCC TAT AGT GA

- complementary oligo: GTG TGC GTA CAA CTA GCT

**Construction with 2 fluorophores.**

- ps32: TCA CTA TAG GGA CAC AAA ACA AAA GAC AAA AAG TGT GCG TAC AAC TAG CT/3BHQ_1/

- fluo2:/5FAM_6/AGC TAG TTG TAC GCA CAC/3BHQ_3/

- fluo3:/5Cy5/TTT TGT CTT TTG TTT TG

Complementary oligos: GTGT GCG TAC AAC TAG CT and CAA AAC AAA AGA CAA AA.

A master mix for each DNA probe (1 µM of ps32 or ps14 DNA and 1 µM of the fluorescent oligonucleotide(s)) was incubated in 10 mM MES pH 6, 50 mM Mg(OAC)₂, 0.1 mM EDTA on a heat block at 95 °C for 5 min. The heat block was then cooled to room temperature (RT) on the bench.

### 2.4. Unwinding assay

In this unwinding assay, in each run, each condition was performed in tetraplicates. Tetraplicates were prepared with a master mix composed of 11 µl of oligo duplex (1 µM), 28 µl of reaction buffer without ATP (10 mM MES pH 6, 50 mM Mg(OAC)₂, 0.1 mM EDTA) and 1 µl of the helicase (diluted in PBS for a final
concentration indicated in each figure). The mix was incubated at RT for 30 min to favour the formation of an enzyme/DNA probe complex. Subsequently, 1 µl of the complementary oligo (100 µM) is added and 9 µl of this master mix were further distributed in 4 different wells of a 96-well plate.

The 96-well plate was placed into the **real time detection thermocycler** (Models BioRad CFX Connect or BioRad CFX Opus 96) programmed at 4 °C for 5 min to obtain a stabilized fluorescence signal (data acquisitions programmed every 20 s). Then, the **real time detection thermocycler** was put on a hold and opened to allow the manual injection of 1 µL of 20 mM ATP to each well of a 96-well plate using a multichannel pipet, while maintaining the temperature at 4 °C. The addition was carefully performed to ensure uniform distribution of ATP across all wells. Following the addition of ATP, the **real time detection thermocycler** program was resumed: first, the fluorescence was monitored at 4 °C for an additional 5 min to confirm the stability of the signal and the inactive state of the enzyme; then the temperature was increased to 20 °C to activate the enzyme and data acquisitions were programmed every 10 s allowing a complete 96plate reading (down to 7s for fewer samples). The temperature's increase between 4 °C and 20 °C has been measured around 3s, meaning that the temperature increase takes less time than monitoring 1 point of the time course. Subsequently, the monitoring was conducted for a duration of 3000 s, a period largely sufficient to achieve fluorescence new equilibrium. Eventually, at the end of the run, a complete dehybridisation may be performed by the manual addition of 1 µl of 10 M NaOH after putting the **real time detection thermocycler** on a hold. After resuming the monitoring at 20 °C for 2 min, the stabilized fluorescence level is used as a reference value to calculate the % of unwinding, for each replicate independently.

## 2.5. Data analysis

Raw data are generated by the software CFXmaestro (Biorad™). For each replicate, the t = 0 of the run is defined as the last measurement done at 4 °C after ATP addition and before the enzyme activation by the temperature increase. Each timepoint is corrected as follows: $Fluo_{t=x} = (Fluo_{measured \ at \ t=x} - Fluo_{measured \ t=0})/Fluo_{t=1}$ corresponding to the first value acquired at 20 °C. The data are processed using a custom-made script developed with R studio (https://gitbio.ens-lyon.fr/LBMC/prio/helicase/). The mean value and standard deviation for each time point are calculated with 3 or 4 replicates per condition. These data are expressed as Relative Fluorescence Intensity. Alternatively, the data can be converted to "% of unwinding" using the external or the internal normalisation procedure that provides an estimation of complete probe unwinding. Each replicate is independently processed prior to computing the mean value and standard deviation for each time point.

# 3. Results and discussion

## 3.1. Principle of the helicase assay in a real time detection thermocycler

To monitor helicase activity of UPF1, we selected two types of DNA probes: Setup-I which was previously utilized [24,30] and Setup-II which we expected to offer higher sensitivity. Setup-I comprises a 18 nt DNA with a 6-FAM fluorochrome at its 5' extremity hybridised to a 50-nt DNA yielding a 32-nt long overhang at the 5' end of the DNA duplex (Fig. 1A). UPF1-HD binds this 5' overhang and translocates 5'-3' on its substrate releasing the fluorescent DNA probe. A complementary 18 nt DNA—bearing a black hole fluorescence quencher (BHQ₁) at its 3' end and present in

excess in the reaction medium—hybridises to the unwound fluorescent DNA resulting in a decrease in fluorescence. In the Setup-II (Fig. 1B), the BHQ₁ at the 3' end of the 50-nt DNA initially quenches fluorescence of the 6-FAM-labelled 18 nt DNA. This combination of blackhole and fluorophore was successfully used in the past for the study of several other helicases [25,31]. UPF1-HD-mediated unwinding causes release of the fluorescent DNA, which then hybridises to a complementary 18 nt unlabelled DNA (present in excess), yielding an increase in fluorescence. The reaction occurs at a controlled temperature (20 °C, except where indicated otherwise) in a **real time detection thermocycler** to monitor fluorescence in real time. We successfully used Qiagen Rotorgene (data not shown), BioRad CFX connect and BioRad CFX opus 96 **real time detection thermocyclers**; data collections were processed using a homemade R script adapted to BioRad specific output files.

## 3.2. Experimental considerations

Our first improvement of a bulk fluorescence-based helicase assay consists in using a real-time detection thermocycler to measure variations in fluorescent signal rather than expensive and unwieldy spectrofluorometers described in the literature. Effectively, real-time detection thermocyclers are compact and cost-effective instruments that enable monitoring in real time fluorescent signal of several wavelengths with high sensitivity and precise control of reaction temperature. However, unlike commonly used spectrofluorometers, real-time detection thermocyclers lack injectors to simultaneously control the initiation of the reaction and monitor the fluorescence baseline, indispensable for a correct interpretation. Notably, ATP can be added manually to initiate the helicase activity at the reaction temperature; nevertheless, by the time the detection thermocycler resumes fluorescent detection, a non-negligible proportion of the oligonucleotide would have already been processed, rendering it impossible to quantify the enzyme activity in its exponential phase. Conversely, if the enzyme is maintained in an inactive state at low temperature, the fluorescence baseline can be evaluated prior to the initiation of the reaction under the requisite conditions. Therefore, our second improvement of the experimental system consists in using temperature as a triggering parameter: DNA duplexes and UPF1-HD are mixed at room temperature and then incubated at 4 °C. Monitoring fluorescence before and after the addition of ATP at 4 °C showed negligible enzymatic activity under these conditions and within a 10-min limit (Fig. 2A and B). However, the Setup-I seemed less accurate as we observed a decrease in fluorescent signal to a greater extent than in the Setup-II, especially at higher enzyme concentrations.

After adding ATP and monitoring signal stability at 4 °C, a temperature increase (up to 20 °C) was programmed and fluorescence levels were measured in real time for at least 3000 s. In Setup-I, the signal exponentially decreases, while in Setup-II it exponentially increases, as expected (Fig. 2C and D). We also observed that the amplitude of the signal was greater in Setup-II than in Setup-I, even though the concentrations of the substrate, ATP and enzyme were the same. Together, the stability of the signal at 4 °C and signal amplitude led us to conclude that Setup-II should be employed for further characterisation of this system.

## 3.3. Qualitative validation of the signal specificity

In order to validate the specificity of the signal obtained with the Setup-II configuration, we used two UPF1-HD inactive mutants UPF1<sub>DE636AA</sub> and UPF1<sub>R843C</sub>. UPF1<sub>DE636AA</sub> has a dual mutation in the ATPase pocket and UPF1<sub>R843C</sub> is a point mutation in the RecA2 domain close to the 5' exit of the RNA binding channel. Both
<figure><img src="image_2.png" /><figcaption>**Fig. 1.** Two experimental setups to monitor the helicase activity of UPF1-HD using fluorescent probes. A) Inspired by Fritz et al., Setup-I follows the extinction of the fluorescence that is being quenched upon unwinding. The upper lane describes the different reaction steps: preparation of UPF1-HD protein (a) and its DNA substrate (b), preloading of UPF1-HD on the DNA substrate (c), unwinding of the fluorescent DNA probe after manually adding ATP (e) and trapping the fluorescent DNA probe with a complementary quencher DNA (f). The middle lane shows the lightcycler program and indicates the theoretical fluorescence emission. The lower lane shows the inactive and active states of UPF1-HD, depending on the temperature and ATP addition. Collected data are processed using custom-made scripts developed with RStudio to generate the normalised graphs of fluorescence as a function of time. B) In Setup-II, the fluorescence is initially quenched; increase in fluorescent signal is detected upon DNA unwinding.</figcaption></figure>

mutants are reported for their ATPase and Helicase deficient activity [30]. As a negative control we heat-inactivated the UPF1-HD<sub>WT</sub>. As anticipated, increase of fluorescence was observed in the UPF1-HD<sub>WT</sub> condition, whereas mutants and heat-inactivated protein were not significantly different from the baseline at both low and high protein concentrations (Fig. 3A and B and Supplementary Fig. 1A). The UPF1 HD enzyme was diluted up to 250 nM and provided a robust helicase activity signal. Previous UPF1 bulk helicase assays were based on lower concentration of UPF1 (~75-50 nM) but performed at 30 °C instead of 20 °C in this manuscript [21,30].

Next, we performed a second control of signal specificity by using the retroviral protein Tax from HTLV-1. We previously showed that Tax inhibits ATPase activity of UPF1 and affects the helicase processivity of UPF1 [29]. As expected, our assay measured decreased helicase activity when UPF1-HD was produced and purified in complex with GST-Tax, thereby demonstrating the capacity of our assay to discriminate between active and inactive UPF1-HD (Fig. 3C and D).

Lastly, since UPF1 cellular activity is mostly described with RNA, we also checked this protocol with an RNA substrate: a 32 nt long RNA coupled with BHQ1 at the 3' extremity was hybridized with the fluorescent DNA probe, mimicking the DNA probe used previously.

The assay was then performed as previously described and we could monitor a specific helicase activity, although the signal was of lower intensity compared with the DNA probe (Fig. 3E).

### 3.4. Quantitative analysis of the helicase assay

To validate the quantitative relevance of our system, we monitored the helicase activity of UPF1-HD with increasing amount of DNA duplexes (Fig. 4A). After 3000 s, the fluorescence levels correlated well with the initial concentration of DNA duplexes (R² = 0.9974, Fig. 4B). Interestingly, the assay is reliable as soon as 55 nM of fluorescent probe, on a similar scale than other fluorescent assays [24,25,30] but at higher concentration than in radio-labelled assays, as expected [21]. We also tried to serial dilute the reaction mix (including the enzyme and the probe) to characterize the minimal conditions of use. As shown in the Supplementary Fig. 1B, dilutions between 1/10 and 1/100 of the conditions described in the method section still allow to obtain a signal with a specific activity compared to conditions without ATP. However, the 1/100 dilution may be too strong to analyse slight variations of the helicase activity.

Furthermore, in order to assess the unwinding efficiency of UPF1-HD in these conditions, it was necessary to measure the
<figure><img src="image_3.png" /><figcaption>**Fig. 2.** Monitoring effect of ATP and low temperature on helicase activity in a real time detection thermocycler. Helicase activity of UPF1-HD was measured at 4 °C (A–B) and at 20 °C (C–D) with or without 2 mM ATP in Setup-I (A, C) and in Setup-II (B, D). To facilitate the comparison between different conditions, fluorescent signal was normalised by its baseline fluorescence at t = 0. Solid lines correspond to mean values from four replicates; light regions displayed in the same colour correspond to standard deviation measured in the four replicates. White lanes indicate a time window of 5min.</figcaption></figure>

<figure><img src="image_4.png" /><figcaption>**Fig. 3.** Characterisation of the assay specificity by helicase inhibition. A) Helicase assay is performed with WT, mutated (DE636AA or R843C) and denaturated UPF1-HD, as indicated. Each graph represents the results obtained with 0.25 µM, 0.5 µM, 1 µM and 2 µM of UPF1-HD. B) Validation of the purity and the levels of the proteins used in A). C) Evaluation of UPF1-HD levels in the UPF1-HD/GST and UPF1-HD/GST-Tax complexes. D) Helicase assay monitored with the complexes UPF1-HD/GST and UPF1-HD/GST-Tax. Reactions deprived of ATP or UPF1-HD are used as negative controls. Solid lines in A and D correspond to mean values from four replicates; light regions displayed in the same colour correspond to standard deviation measured in the four replicates. E) Helicase assay was monitored with a RNA probe. As a comparison, the assay was run in parallel with a DNA probe. Both probes hybridized with the same fluorescent DNA oligo. Negative controls were performed without ATP ("-ATP").</figcaption></figure>

maximum fluorescence signal achievable for each condition (% unwinding). To do so, we proposed two different methods: (1) an external normalisation that relies on measuring fluorescent signal of the fluorescent oligonucleotide pre-hybridised with the trapping oligonucleotide, prior to adding the long quenching oligonucleotide

and the enzyme. By doing so, we mimic the result of a 100 % efficient unwinding reaction. Monitoring the dose response of fluorescent oligonucleotides prepared under these conditions revealed a strict linear signal between 140 nM and 965 nM, indicating the effective range of use (Supplementary Fig. 2A); (2) an internal
<figure><img src="image_5.png" /><figcaption>**Fig. 4.** Quantitative characterisation of the helicase assay. A) Monitoring the helicase assay with a fixed concentration of UPF1 (1 µM) and increasing amount of DNA substrate through relative fluorescence intensity. B) The maximal fluorescence intensity from A (obtained at 3000 s) is represented as a function of DNA duplex concentration. C) Data from the helicase assay in A are represented as a % of the total DNA duplex unwinding. Data were fit to $f(t) = A \times (1 - e^{-k_1 \times t}) + k_2 \times t$, where the constants A, k₁ and k₂ are indicated for each condition in the lower part. D) same as C) with increasing concentrations of UPF1-HD and 275 nM DNA probes. E) same as C, with increasing concentrations of ATP and 0.25 µM UPF1-HD). F) same as C) with increasing temperature and 0.5 µM UPF1-HD. Solid lines in A, C and D–F correspond to mean values from four replicates; light regions displayed in the same colour correspond to standard deviation measured in the four replicates.</figcaption></figure>

normalisation, that relies on dehybridisation of the persisting DNA duplexes at the end of each experiment by adding 1 M (final concentration) NaOH (Supplementary Fig. 2B). An internal normalisation has the advantage to be performed from each monitored sample; however, the basification of the reaction medium may also affect fluorescent probe emission [32]; therefore, the method of normalisation may be specific of each experimental protocol and has to be investigated beforehand. In our setup, both normalisation approaches yielded similar evaluation of duplexes unwinding percentage (Supplementary Figs. 2A–B).

Applying the external normalisation method to monitor UPF1 activity with increasing amount of DNA duplexes revealed that the concentration of the DNA probe does not affect the percentage of unwinding to the contrary of the Relative Fluorescence Intensity

evaluation (compare Fig. 3A and C). This was expected since UPF1-HD was preloaded on the DNA duplexes before the reaction initiation (Fig. 3C). By contrast, unwinding efficiency increased with increasing amounts of enzyme and ATP or with higher temperature (Fig. 3D–F).

Furthermore, in these conditions, the real-time emission of fluorescence fitted with the equation $f(t) = A \times (1 - e^{-k_1 \times t}) + k_2 \times t$ in which A and k₁ represent the amplitude and the rate constant of the exponential burst phase and k₂ is the rate constant of the linear steady-state phase. This equation has been previously shown to be an adequate model of helicase activity [33], confirming that our assay is quantitatively relevant. k₂ is due to some reassociation of UPF1-HD with a new substrate after unwinding, even though the trapping oligonucleotide was present
in excess. Addition of heparin may reduce this multiple turnover [20]. Therefore, we added heparin in the reaction mix, what successfully prevented UPF1-HD multiple turnover without altering its helicase activity (Supplementary Fig. 3). Interestingly, our kinetic also analysis revealed that optimal ATP concentrations were not the same to obtain a helicase activity with a maximal amplitude (A) or a maximal initial burst of reaction ($k_1$) (Fig. 3E and Supplementary Fig. 4). As indicated by the profile of amplitude, we found that in our experimental conditions, the optimum of ATP is around 20 µM. To the opposite, when analysing $k_1$, 2 mM ATP allow an intense burst of activity as soon as the temperature increases. However, the decrease of amplitude in these conditions clearly implies that such an activity is not adapted for UPF1 on the longer term, certainly leading to discontinued unwinding events. A different profile may be observed with other helicases and it would deserve further investigations.

### 3.5. Impact of length and position of double-stranded region on UPF1-HD helicase activity

To illustrate the potential of our approach to reveal subtle differences in helicase efficiencies, we investigated the impact of the length of the double-stranded region of the probe on UPF1-HD unwinding kinetics. In our initial Setup-II, the DNA probe (i) consists of a 18 nt double-stranded region and a 32 nt single-stranded overhang at the 5' end and enables binding of up to two UPF1-HP molecules (Fig. 5A). To extend the duplex length, we hybridised an 18 nt, unlabelled DNA oligonucleotide upstream of the double-stranded region (Fig. 5A probe (ii)); in this configuration essentially one UPF1-HD molecule can bind the remaining 14 nt long overhang. We also designed two additional DNA probes: probe (iii) has a short duplex and a short overhang and probe (iv) is 50 nt long, double-stranded dsDNA with a central, 18 nt single-strand region (Fig. 5A); both probes (iii) and (iv) can be bound by one UPF1-HD molecule upstream of a short DNA duplex to be unwound.

Gel-shift experiments showed similar amounts of UPF1-HD loaded onto all four probes (Fig. 5B). Interestingly, UPF1 unwound the probe (ii) slower than the probe (i). This decreased overall efficiency of helicase activity is not due to longer double-stranded region of probe (ii), because we observed the same kinetics for probe (iii) consisting of 18 nt long DNA duplex. Moreover, the probe (iv) that differentiates from the probe (iii) by the presence of a dsDNA at the 5' side of UPF1-HD, displayed the same kinetics as the probe (i) (Fig. 5C). Thus, these data indicated that UPF1 is more efficient at unwinding when it is not bound to the extremity of the probe.

### 3.6. Fluorescent dual-probe method for measuring helicase activity

The detection thermocycler can monitor multiple fluorophores in the same run to achieve multiplexing. Therefore, we sought to extend our method to quantitative analysis of helicase activity using two adjacent probes labelled in different wavelengths. We hybridised the long BHQ₁ ssDNA to two short DNA molecules labelled with Cy5 or FAM-6 on their 5' end (Fig. 5D). The FAM-6 fluorescence is quenched by the BHQ₁ at the 3' end of the complementary long ssDNA while the Cy5 fluorescence is quenched by a BHQ₃ at the 3' end of the FAM-6 labelled oligo, as shown in Fig. 5D. We confirmed the absence of fluorescence leakage between the 2 channels (Fig. 5E). Monitoring helicase activity of UPF1-HD yielding overlaying graphs, suggesting that both oligonucleotides were unwound with the same kinetics (Fig. 5E). We did not observe a time delay between unwinding of the two DNA molecules; this time delay is likely shorter than the minimal time that the detection thermocycler requires to sequentially read fluorescent signal of

each channel in real-time. Nevertheless, we expect that using longer probes will overcome this technical limitation and will enable measuring the mean speed of helicases unwinding nucleotide duplexes. Of note, the Cy5 emission yields stronger signals than FAM-6 (Fig. 5E, Relative Fluorescence Intensity), suggesting a potential for enhanced sensitivity in this assay, beneficial for less efficient helicases.

## 4. Conclusion

Here, we sought to improve existing in vitro bulk measurement of helicase activity. Considering strong limitations associated with the use of radioactivity and expensive cost of the real-time plate reader spectrofluorometers combined with injectors, we thought of using a real-time detection thermocycler and described in this report the specific experimental setup required to control the initiation of helicase activity and to monitor unwinding with temperature control. We benchmarked our method using UPF1-HD and demonstrated that our method yields quantitative measurements of helicase activity and is able of revealing subtle differences in unwinding of different substrates. Most of the helicases have an optimal activity above 20 °C and are considered to exert a reduced activity at 4 °C. This also include helicases from hyperthermophilic species. As such, they should be qualified for this assay. However, some specific helicases have been shown to play an important role in the adaptation to cold chock [34] or in psychrotolerant organisms (review in Refs. [35,36]). In these cases, we may expect them to work at low temperature (at which their expression is induced) and thus not be fitted for this assay, unless specific adaptations.

Recently, new microplate reader solutions with customised options including injectors, thermal control and fluorescence filters were available at a more affordable price. Adaptation of our protocol to these technologies should be possible. However, the use of a thermocycler as described here still have several advantages: it provides a quick and precise control of the temperature from 4 °C to at least 70 °C what may be critical for hyperthermophilic helicases; thermocyclers are already widely widespread in labs including those not specialized in fluorescent monitoring assays; finally, thermocyclers allow the simultaneous monitoring of several fluorophores in real time. Therefore, our method can monitor unwinding of several probes labelled with different fluorophores, enabling studies of helicase directional activity ($5' \rightarrow 3'$ or $3' \rightarrow 5'$), the impact of secondary structures on the helicase activity, as well as measurement of the mean speed of the helicase of interest. Last, but not least, our assay runs in a 96-well plate format and requires a reduced reaction volume, thereby enabling semi high-throughput route for defining molecular mechanisms of helicases.

### CRediT authorship contribution statement

**Jean-Philippe Robin:** Writing – original draft, Visualization, Software, Investigation, Formal analysis, Data curation, Conceptualization. **Vincent Mocquet:** Writing – original draft, Supervision, Methodology, Funding acquisition, Conceptualization.

### Data availability statement

R scripts are available at https://gitbio.ens-lyon.fr/LBMC/prio/helicase/.

### Funding sources

This work was supported by a "projet émergent grant" to VM from the Ecole Normale Supérieure de Lyon (ENSL).
<figure><img src="image_6.png" /><figcaption>**Fig. 5.** Revealing differences of helicase activity using different substrates A) Schematic representation of different DNA substrates used in this study. B) EMSA carried out after UPF1-HD UV crosslinking on each substrate. C) Monitoring helicase assay using each substrate. D) Schematic representation of the helicase assay using two sequential fluorescent probes (Cy5 and FAM-6 are represented by red and green stars). E) Monitoring helicase assay with the two emission channels (670 nm for Cy5 and 520 nm for FAM-6). The absence of FAM-6 signal at 670 nm and Cy5 at 520 nm was controlled with the indicated substrates (right panel). Superposition of the fluorescent signals measured in the 670 nm and 520 nm channels (left part). Solid lines in C and D correspond to mean values from four replicates; light regions displayed in the same colour correspond to standard deviation measured in the four replicates.</figcaption></figure>

**Declaration of competing interest**

None.

**Acknowledgements**

The authors thank Stephane Réty, Pierre Jalinot, Karina Jouravleva and Hélène Polvèche for critical discussions and corrections.

**Appendix A. Supplementary data**

Supplementary data to this article can be found online at
https://doi.org/10.1016/j.biochi.2025.02.002.

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