Assessment of Tfh Cells and Related Molecules in CML‐CP Patients With Different Molecular Responses

ABSTRACT

Introduction

This study aimed to characterize T follicular helper (Tfh) cell differentiation and related molecular profiles in chronic‐phase chronic myeloid leukemia (CML‐CP) patients with distinct molecular responses and to explore immune correlates of treatment outcomes.

Methods

Peripheral blood CD4⁺ T cells were analyzed in 125 CML‐CP patients (8 newly diagnosed, 71 MMR, and 46 pre‐MMR) and 22 healthy donors. We assessed CD4⁺ T subset distribution and expression of Tfh‐related molecules including Bcl6, Blimp1, PD‐1, PD‐L1, and PD‐L2.

Results

Newly diagnosed and pre‐MMR patients exhibited decreased CD4⁺ T cell proportions compared to healthy donors and MMR patients (p < 0.05), while pre‐MMR patients had elevated frequencies of CD4⁺CXCR5⁺PD‐1⁺ Tfh cells and increased PD‐1 expression on CD4⁺ T cells (p < 0.05). Bcl6, PD‐1, and PD‐L1 were upregulated in pre‐MMR cases, whereas Blimp1 and PD‐L2 remained unaltered.

Conclusion

Pre‐MMR CML‐CP is accompanied by aberrant Tfh cell differentiation and an immunosuppressive immune signature linked to suboptimal molecular responses. Targeting immune checkpoints or modulating Tfh cell homeostasis may help improve therapeutic outcomes in CML‐CP.

1. Introduction

Chronic myeloid leukemia (CML) is a malignant tumor caused by clonal proliferation of hematopoietic stem cells in the bone marrow, accounting for 15% of adult leukemia's. It can occur in all age groups (SEER statistics), with a median age at diagnosis of 67 years [1]. However, CML patients tend to be younger in China, with a median age of 45–50 years [2]. The global annual incidence is 1.6–2 per 100 000 population [3]. The hallmark genetic alteration in CML is the t(9;22)(q34;q11) chromosomal translocation (known as the Philadelphia chromosome), leading to the formation of the BCR‐ABL1 fusion gene. This gene encodes an aberrant tyrosine kinase protein that drives malignant proliferation of myeloid cells through sustained activation of downstream signaling pathways [4].

With the application of first‐generation tyrosine kinase inhibitors (TKIs), the 10‐year survival rate of CML patients has significantly increased to 85%–90% [5]. Second‐generation TKIs used as first‐line therapy enable faster and deeper molecular responses [6, 7]. However, some patients remain insensitive to TKI treatment. According to the cancer statistics 2023, an estimated 8930 people will be diagnosed with CML in the United States, and 1310 people will die of the disease [8], with a mortality rate as high as 14.7%. Therefore, improving TKI sensitivity in CML patients or developing better therapeutic approaches to help TKI‐insensitive CML patients achieve major molecular response (MMR), deep molecular response (DMR), and treatment‐free remission (TFR) remains critically important.

Increased studies suggested that the efficacy of TKI in the treatment of CML patients is affected by host immune system, especially in achievement of DMR to promote TFR and improve the quality of life. T cell‐mediated immune responses can play an important role in the regulation of CML cells and immune microenvironment, such as Treg‐mediated immune suppression [9, 10]. The differentiation proportions of Tregs decreased in CML‐CP patients with TKIs therapy, especially patients with DMR [11]. Tfh cells were an important subset of CD4⁺ T cells, which characteristically expressed CXCR5, PD‐1, Bcl‐6 and other molecules [12]. Tfh cells have both anti‐tumor and pro‐tumor roles in tumor immunity [13, 14]. However, the roles of Tfh cells in CML patients on therapy remain unknown.

In this study, we evaluated the differentiation characteristics of CD4⁺ T cells and Tfh cells in patients with CML under different molecular responses, as well as the expression levels of the key transcription factor Bcl‐6 and the inhibitory molecules PD‐1, PD‐L1, and PD‐L2. By analyzing the phenotypic evolution pattern of Tfh cell subsets during the process of molecular remission, this research focused on exploring the possible immunoregulatory function of Tfh cells on the achievement of MMR and DMR in CML patients with TKIs therapy. The results of this study will provide a new therapeutic strategy on targeting immune checkpoints or modulating Tfh cell differentiation to promote the CML patients on TKIs therapy to achieve MMR and DMR.

2. Methods

2.1. Patients and Sample Collection

Peripheral blood samples were collected from 125 CML‐CP patients with different molecular responses on TKIs therapy and 22 healthy donors. Among these patients, 8 were newly diagnosed and sampled prior to any TKI treatment. The remaining 117 patients (pre‐MMR and MMR groups) were sampled after at least 12 months of continuous TKI therapy, aligning with the treatment milestones recommended by the European LeukemiaNet (ELN) guidelines. Baseline characteristics, including age, sex, and the type and duration of TKI therapy, were collected and are summarized in Table 1. This study was approved by the ethics Committee of Guangdong Provincial People's Hospital (KY‐Q‐2021‐284‐01) and conducted in accordance with the Declaration of Helsinki. BCR‐ABL1 fusion gene levels have been converted to international standardized values and are expressed as percentages.

TABLE 1.

Baseline characteristics of 125 CML‐CP patients and 22 healthy donors.

Characteristic	New diagnosis (n = 8)	Pre‐MMR (n = 46)	MMR (n = 71)	Healthy donors (n = 22)
BCR‐ABL1 (IS)	≥ 10%	> 0.1%	≤ 0.1%	0
Age (y), median (range)	38.5 (26–65)	35 (10–63)	39 (17–80)	26 (22–45)
Gender, n (%)
Male	3 (37.5%)	26 (56.5%)	39 (54.9%)	14 (63.6%)
Female	5 (62.5%)	20 (43.5%)	32 (45.1%)	8 (36.4%)
TKI type, n (%)
Imatinib	0	33 (71.7%)	59 (83.1%)	—
Nilotinib	0	6 (13.1%)	8 (11.3%)	—
Dasatinib	0	7 (15.2%)	4 (5.6%)	—
TKI Duration (mo), Median (range)	0	21.27 (12.1–132.63)	41.47 (12.13–151.07)	—

2.2. Real‐Time PCR Analysis

Total RNA was extracted from peripheral blood cells using TRIzol reagent (Invitrogen, USA), and its purity (A260/A280 ratio 1.8–2.0) and concentration were determined using a NanoDrop 2000 spectrophotometer (Thermo Scientific, USA). One microgram of total RNA was reverse transcribed into cDNA using the PrimeScript RT kit (Takara, Japan) under the following reaction conditions: 30 min at 37°C and 5 s at 85°C. An absolute quantification method based on a standard curve was employed. Known concentrations of ABL were used as standards to construct standard curves through serial dilutions. The ABL reference gene, BCR‐ABL1 (P210) fusion gene, and Tfh‐related target genes (Bcl6, Blimp1, PD‐1, PD‐L1, and PD‐L2) were quantified using this standard curve approach. The absolute copy numbers or relative quantities of each target gene were calculated by interpolating from their respective standard curves. The expression level of each target gene was then normalized to the ABL reference gene by calculating the ratio of target gene quantity to ABL quantity. The primers and probe sequences were designed by Primer Express and synthesized by BGI (Shenzhen) after specificity verification by BLAST. The primer sequences are shown in Table 2. Amplification efficiency was verified to be 90%–110% via standard curves (see also Figure S1).

TABLE 2.

Primer lists.

Primer name	Sequence (5′‐3′)
ABL‐F	TGGAGATAACACTCTAAGCATAACTAAAGGT
ABL‐R	GATGTAGTTGCTTGGGACCCA
ABL‐P	CCATTTTTGGTTTGGGCTTCACACCATT
P210‐F	TCCGCTGACCATCAAYAAGGA
P210‐R	CACTCAGACCCTGAGGCTCAA
P210‐P	CCCTTCAGCGGCCAGTAGCATCTGA
Bcl6‐F	GCCGGCTGACAGCTGTATC
Bcl6‐R	CGGAGACGATTAAGGTTGAGAAG
Bcl6‐P	AGTTCACCCGCCATGCCAGTGATG
PD‐1‐F	GGATTTCCAGTGGCGAGAGA
PD‐1‐R	GGAAAGACAATGGTGGCATACTC
PD‐1‐P	CCCCCCGTGCCCTGTGTCC
Blimp1‐F	CGCCTTGAGCATGAAGCA
Blimp1‐R	GTCCTTGTTCTGTGGGTACTTGACT
Blimp1‐P	AGGCCAGCTCCACCGATGACAAGAA
PD‐L1‐F	AAAGAATTTTGGTTGTGGATCCA
PD‐L1‐R	TCACTGCTTGTCCAGATGACTTC
PD‐L1‐P	ACATGTCAGGCTGAGGGCTACCCCA
PD‐L2‐F	CATCCAACTTGGCTGCTTCA
PD‐L2‐R	TTAGGGCTATCACTGTGGCTATGA
PD‐L2‐P	CATCCCCTTCTGCATCATTGCTTTCATT

2.3. Flow Cytometry Analysis

EDTA‐anticoagulated peripheral blood was collected from CML‐CP patients, and red blood cells were removed using red blood cell lysis buffer to isolate peripheral nucleated cells. For cell surface staining, 1 × 10⁶ cells were suspended in staining buffer and incubated with fluorescein‐labeled antibody mixture containing anti‐CD4‐FITC, anti‐CXCR5‐APC, and anti‐PD‐1‐PE for 30 min at room temperature (25°C) in the dark. Isotype antibodies and fluorescence‐minus‐one (FMO) controls were used for gating correction. After staining, cells were washed twice with phosphate‐buffered saline (PBS) and resuspended in PBS containing 1% fetal bovine serum (FBS). Data were acquired using a BD FACSCanto II flow cytometer equipped with 488 nm and 640 nm lasers; at least 10 000 lymphocyte events were collected per sample. FlowJo v10.8 software was used for data analysis. The sequential gating strategy was as follows (see also Figure S2):

(A) Lymphocytes were first gated based on forward scatter (FSC) and side scatter (SSC) to exclude debris and dead cells, (B) CD4⁺ T cells were identified from the lymphocyte population, (C) PD‐1 expression on CD4⁺ T cells was analyzed within the CD4⁺ gate, (D) CD4⁺CXCR5⁺PD‐1⁺ double‐positive cells were defined as the T follicular helper (Tfh) cell subset. This gating strategy was applied consistently across all samples. The definition of Tfh cells as CD4⁺CXCR5⁺PD‐1⁺ is widely accepted for identifying circulating Tfh (cTfh) cells in human peripheral blood, as CXCR5 directs migration to B cell follicles and PD‐1 serves as a reliable activation/maturation marker [15, 16]. Intracellular Bcl‐6, while lineage‐defining, is not suitable for live‐cell surface staining in routine flow cytometry. Therefore, the combination of CXCR5 and PD‐1 remains the standard surrogate for enumerating cTfh cells in human studies.

2.4. Statistical Analysis

Experimental data were expressed as mean ± standard deviation (mean ± SD) or median (interquartile range) [median (IQR)], depending on the data distribution. Normality was assessed using the Shapiro–Wilk test, and homogeneity of variances was evaluated using Levene's test. The normality testing results are summarized in Table S1. For variables that did not follow a normal distribution (e.g., Tfh proportions, PD‐1, and Bcl6 expression), non‐parametric tests (Mann–Whitney U) were used. For normally distributed variables (e.g., CD4⁺ T cell percentages, BLIMP1, PD‐L1, and PD‐L2 expression), parametric tests (t‐test or ANOVA) were applied. The comparison strategy was as follows: For two independent groups, Student's t‐test was used when data followed a normal distribution and variances were equal; otherwise, the Mann–Whitney U test was applied. For three or more independent groups, one‐way analysis of variance (ANOVA) was performed when normality and homogeneity of variances assumptions were met. Following a significant ANOVA result, post hoc multiple comparisons were conducted using Tukey's honestly significant difference (HSD) test, which provides a more conservative adjustment for the family‐wise error rate. When the assumptions of normality or homogeneity of variances were violated, the Kruskal–Wallis test was used, followed by post hoc test with Bonferroni correction for multiple comparisons. All tests were two‐sided, and a significance threshold of α = 0.05 was applied. All statistical analyses were performed using GraphPad Prism v9.0 and SPSS v27.

3. Results

3.1. The Proportion of CD4 ⁺ T Cells in CML‐CP Patients on TKIs Therapy With Different Molecular Responses

Flow cytometric analysis was performed to detect the proportion of CD4⁺ T cells in peripheral blood lymphocytes, with the representative flow cytometry gating strategy shown in Figure 1A. CD4⁺ T cell proportions were observed among CML‐CP patients receiving TKI therapy in different molecular response stages (Figure 1B). Compared with newly diagnosed CML‐CP patients (13.40% ± 6.25%), both the pre‐MMR group (30.19% ± 7.24%) and MMR group (36.16% ± 8.31%) exhibited significantly higher CD4⁺ T cell proportions (all p < 0.001). The percentage of CD4⁺ T cells in the MMR group was comparable to that in healthy donors (35.54% ± 7.89%), and was significantly higher than that in pre‐MMR patients (p < 0.001). Pre‐MMR patients showed intermediate CD4⁺ T cell levels, which were markedly elevated relative to newly diagnosed cases (p < 0.001). These findings suggest a progressive restoration of CD4⁺ T cell immunity correlating with improved molecular responses during TKIs treatment, highlighting the potential immunological benefits associated with achieving deeper molecular remission (DMR) in CML‐CP management.

FIGURE 1.

Frequency of CD4⁺ T cells within lymphocytes in CML‐CP patients with different molecular responses. (A) Representative flow cytometry gating strategy for identifying CD4⁺ T cells. (B) Comparison of the percentages of CD4⁺ T cells among four groups are as follows: Healthy donors (35.54% ± 7.89%), newly diagnosed CML‐CP patients (13.40% ± 6.25%), pre‐MMR patients (30.19% ± 7.24%), and MMR patients (36.16% ± 8.31%). One‐way ANOVA was performed for intergroup comparison.

3.2. The Expression of PD‐1 on CD4 ⁺ T Cells Decreased in MMR CML‐CP Patients

Analysis of PD‐1 expression on CD4⁺ T cells revealed obvious differences among CML‐CP patients at different molecular response stages. The flow cytometry gating strategy for detecting PD‐1 expression on CD4⁺ T cells is shown in Figure 2A. As presented in Figure 2B, PD‐1 expression levels were compared across four groups: healthy donors (5.79% [3.58%], n = 22), newly diagnosed CML‐CP patients (15.33% [11.88%], n = 8), pre‐MMR patients (11.80% [14.26%], n = 46), and MMR patients (6.71% [5.79%], n = 71). MMR patients showed PD‐1 expression comparable to healthy donors, while both groups exhibited significantly lower PD‐1 levels than pre‐MMR individuals. Newly diagnosed patients displayed an intermediate PD‐1 expression profile, with no statistically significant differences versus the other subgroups. Such lack of statistical difference may be attributed to the relatively small sample size of the newly diagnosed cohort. These findings indicate that attainment of molecular remission under TKI treatment is accompanied by normalized PD‐1 expression on CD4⁺ T cells. In contrast, patients with incomplete molecular response (pre‐MMR) maintain elevated PD‐1 levels, implying persistent immune dysregulation.

FIGURE 2.

PD‐1 expression on CD4⁺ T cells in CML patients with different molecular response. (A) PD‐1 expression on CD4⁺ T cell flow cytometry gating strategy. (B) Comparison of PD‐1 expression on CD4⁺ T cells among healthy donors (5.79% [3.58%], n = 22), newly diagnosed CML‐CP patients (15.33% [11.88%], n = 8), pre‐MMR patients (11.80% [14.26%], n = 46), and MMR patients (6.71% [5.79%], n = 71).

3.3. Increased Frequency of Tfh Cells in Pre‐MMR CML‐CP Patients

This study provides the first investigation into the treatment response‐associated dynamic alterations of Tfh cells during TKI therapy for CML‐CP. CD4⁺CXCR5⁺PD‐1⁺ Tfh cells were identified using the flow cytometry gating strategy shown in Figure 3A. A comparison of CD4⁺CXCR5⁺PD‐1⁺ Tfh cell differentiation proportions among the four groups revealed distinct dynamic patterns (Figure 3). Specifically, pre‐MMR patients (n = 46) exhibited significantly increased peripheral Tfh cell proportions (median 3.81% [IQR 2.69%]) compared to both MMR patients (n = 71; 3.02% [1.57%], p < 0.05) and healthy donors (n = 22; 1.49% [1.49%], p < 0.001). Newly diagnosed CML‐CP patients (n = 8) showed intermediate Tfh cell proportions (2.73% [1.86%]), which were not statistically different from those of the other three clinical subgroups (all p > 0.05). Additionally, MMR patients maintained significantly higher Tfh cell levels than healthy controls (p < 0.001). This dynamic pattern suggests that Tfh cell expansion may reflect disease‐specific immune activation during incomplete molecular response, with partial normalization upon achieving MMR. These findings indicate stage‐specific regulatory roles of Tfh cells in CML immunomodulation, which may potentially participate in immune microenvironment remodeling.

FIGURE 3.

Proportion of CD4⁺CXCR5⁺PD‐1⁺ Tfh cells differentiation in CML patients with different molecular response. (A) CD4⁺CXCR5⁺PD‐1⁺ Tfh cells flow cytometry gating strategy. (B) Comparison of CD4⁺CXCR5⁺PD‐1⁺ Tfh cells differentiation proportions among healthy donors (1.49% [1.49%], n = 22), newly diagnosed CML‐CP patients (2.73% [1.86%], n = 8), pre‐MMR patients (median 3.81% [2.69%], n = 46), and MMR patients (3.02% [1.57%], n = 71).

3.4. MMR CML‐CP Patients Have Reduced Tfh‐Related Transcription Factor Bcl6 and PD‐1 Inhibitory Molecule

A total of 83 out of 125 enrolled CML‐CP patients had eligible residual RNA samples for subsequent Tfh‐related molecular analysis, including 33 pre‐MMR and 50 MMR patients. Quantitative real‐time PCR (qRT‐PCR) was performed to detect the peripheral blood expression of Tfh‐associated molecules between the two groups (Figure 4). For molecules presented as median (IQR), PD‐1 expression was significantly lower in MMR patients than in pre‐MMR patients (26.39% [16.89%] vs. 50.44% [26.02%], p < 0.001), and the Tfh lineage‐specific transcription factor Bcl6 was also markedly downregulated in the MMR group (2005.95% [1680.55%] vs. 2969.75% [2446.21%], p < 0.001) (Figure 4A,D). For molecules presented as mean ± standard deviation, MMR patients showed significantly reduced PD‐L1 mRNA levels compared with pre‐MMR patients (58.05% ± 72.30% vs. 105.24% ± 123.93%, p = 0.031). In contrast, no significant intergroup differences were observed in PD‐L2 (10.92% ± 9.73% vs. 15.38% ± 17.64%, p = 0.142) and Blimp1 expression (373.57% ± 154.64% vs. 434.73% ± 146.89%, p = 0.076) (Figure 4B,C,E). These results indicate a selective downregulation of the PD‐1/PD‐L1‐Bcl6 axis in MMR patients, while Blimp1 and PD‐L2 remained relatively unchanged, suggesting targeted modulation of Tfh‐related immune checkpoint molecules upon achieving optimal molecular remission during TKI therapy.

FIGURE 4.

The expression of Tfh‐related molecule in peripheral blood cells in CML patients on TKI therapy. (A) Comparison of the expression of PD‐1 inhibitory molecule between pre‐MMR (50.44% [26.02%], n = 33) and MMR CML‐CP patients (26.39% [16.89%], n = 50), p < 0.001. (B) The expression of PD‐L1 in the peripheral blood of MMR (58.05% ± 72.30%, n = 33) and pre‐MMR CML‐CP patients (105.24% ± 123.93%, n = 50) was compared, p = 0.031. (C) Comparison of the expression of PD‐L2 between pre‐MMR (15.38% ± 17.64%, n = 33) and MMR (10.92% ± 9.73%, n = 50) CML‐CP patients, p = 0.142. (D) Comparison of the expression of Bcl6, a key transcription factor of Tfh, between pre‐MMR (2969.75% [2446.21%], n = 33) and MMR CML‐CP patients (2005.95% [1680.55%], n = 50), p < 0.001. (E) Comparison of the expression of Blimp1 between pre‐MMR (434.73% ± 146.89%, n = 33) and MMR (373.57% ± 154.64%, n = 50) CML‐CP patients, p = 0.076.

4. Discussion

CD4⁺ T cells, as central regulators of the immune system, play multifaceted roles in anti‐tumor immunity and demonstrate multidimensional effects in hematologic tumor immunity—ranging from direct tumor killing to immune remodeling [17, 18, 19]. While traditionally considered to function indirectly through assisting CD8⁺ T cells [20], recent studies [21, 22, 23, 24, 25] have revealed their capacity to directly eliminate tumor cells via mechanisms such as secreting IFN‐γ and TNF‐α to induce tumor cell apoptosis or cell cycle arrest, overexpressing IL‐2, differentiating into cytotoxic T lymphocytes (CTLs), or activating macrophages to release inflammatory mediators like nitric oxide for indirect tumor cell elimination. Additionally, they exert anti‐tumor immune effects by remodeling the immune microenvironment [26, 27].

During CML progression and imatinib treatment, the role of the PD‐1/PD‐L1 pathway, which was reported as a critical immune checkpoint mechanism mediating immune escape, T‐cell exhaustion, and differential therapeutic responses [28]. Leukemic cells and myeloid cells in the CML microenvironment upregulate PD‐L1 expression, while CD4⁺ and CD8⁺ T cells display elevated PD‐1 levels. This generates inhibitory PD‐1/PD‐L1 signaling that directly suppresses antitumor T‐cell activity, drives localized immune exhaustion, and supports leukemic immune escape and expansion [29]. In early treatment, PD‐1/PD‐L1 signaling predominates in non‐optimal responders [30]. PD‐L1 blockade fails to restore T‐cell proliferation but upregulates IL‐2 secretion, confirming its involvement in T‐cell dysfunction [31]. Dynamic changes in PD‐1/PD‐L1 are closely associated with CML risk stratification, bone marrow microenvironment remodeling, and the switch of immunosuppressive networks, positioning this pathway as a key determinant of the efficacy of immunotherapy combined with TKIs.

As a critical subset of CD4⁺ T cells, Tfh cells—characteristically expressing CXCR5, PD‐1 and Bcl‐6—act as a “double‐edged sword” in tumor immunity [32, 33]. While capable of supporting anti‐tumor humoral immunity, they may also be modulated by the tumor microenvironment (TME) in ways that associate with immune escape. In anti‐tumor immunity, Tfh cells drive humoral immune responses by activating B cells to produce tumor‐specific antibodies and enhance CD8⁺ T‐cell function via IL‐21 secretion, thereby promoting effector T‐cell infiltration and cytotoxic activity in the TME and synergizing with cellular immunity [34, 35]. Conversely, Tfh cells have been linked to tumor immune evasion. Proposed mechanisms include: (1) promoting the expansion of regulatory B cell subsets and upregulating IL‐10 and TGFβ secretion to suppress anti‐tumor immunity [36]; (2) associating with T‐cell exhaustion through high expression of inhibitory receptors such as PD‐1 [14]; and (3) recruiting regulatory T cells (Tregs) via CXCL13 to support immunosuppressive microenvironments [37].

In this study, we systematically delineated the immunological landscape of CML‐CP patients under TKI therapy, revealing three critical axes of immune remodeling associated with the depth of molecular response are as follows: (1) quantitative restoration of CD4⁺ T cell populations; (2) qualitative normalization of immune checkpoint expression, especially the PD‐1/PD‐L1 pathway; and (3) stage‐specific alterations in Tfh cell biology. These findings collectively indicate that molecular remission achieved through TKIs is accompanied by multi‐dimensional immune reconstitution, rather than merely a quantitative reduction in BCR‐ABL transcripts.

The progressive recovery of CD4⁺ T cell proportions from newly diagnosed (13.40%) through pre‐MMR (30.19%)–MMR (36.16%) stages (Figure 1) parallels the trajectory of molecular response, supporting the notion that CD4⁺ reconstitution may serve as a potential immunological correlate of treatment efficacy. Notably, CD4⁺ levels in the MMR group were comparable to those in healthy donors (35.54%, p > 0.05), consistent with immune normalization rather than compensatory expansion. This interpretation is supported by parallel PD‐1 expression on CD4⁺ T cells dynamics: while pre‐MMR patients exhibited elevated PD‐1 levels (11.80% [IQR 14.26%], Figure 2), MMR patients displayed PD‐1 levels similar to healthy controls (6.71% [IQR 5.79%] vs 5.79% [IQR 3.58%], p > 0.05), consistent with the resolution of chronic antigen‐driven T‐cell exhaustion.

In addition, we are the first to report that the proportion of circulating Tfh cells in pre‐MMR patients (3.81% [IQR 2.69%]) is significantly higher than in the MMR group (3.02% [1.57%], p < 0.05) and healthy donors (1.49% [1.49%], p < 0.001) (Figure 3). This increase was accompanied by higher expression of Bcl6 and PD‐1/PD‐L1 (Figure 4). These correlative observations suggest that Tfh cell expansion may be associated with an immunosuppressive milieu in pre‐MMR patients. These patterns indicate that incomplete molecular remission is associated with Tfh cell expansion and persistent immune activation, which may coincide with an immunosuppressive state. Further mechanistic studies are needed to explore the functional role of the CXCR5‐PD‐1 axis during TKI treatment in CML. The Tfh levels in the MMR group remained higher than in healthy donors (p < 0.001), which may correlate with sustained immune activation driven by residual leukemic antigens. The clinical significance of this observation requires evaluation in combination with long‐term follow‐up data.

The selective downregulation of the PD‐1/PDL1‐Bcl6 axis in MMR patients (Figure 4), in contrast to stable Blimp1/PDL2 expression, suggests that TKI‐mediated immune changes are associated with specific modulation of the Tfh‐immune checkpoint module rather than global transcriptional reprogramming, which may contribute to immune microenvironment remodeling. This molecular signature—reduced PD‐1/PDL1/Bcl6 with preserved Blimp1—may potentially reflect an immunological state that balances leukemia surveillance and immune homeostasis.

5. Conclusion

In conclusion, this study demonstrates that the recovery of CD4⁺ T cell populations and normalization of PD‐1 expression parallel the depth of molecular remission in CML‐CP patients receiving TKI therapy. These findings validate immune parameters as complementary biomarkers to BCR‐ABL transcript monitoring for assessing treatment response. Additionally, our data suggest that Tfh cell hyperactivity and dysregulated Bcl6 expression may be associated with immune dysfunction, which may impede pre‐MMR patients from achieving MMR. Collectively, combining TKIs with targeted therapies against the PD‐1 immune checkpoint or strategies modulating Tfh cell differentiation holds promise as an important therapeutic approach to help pre‐MMR CML‐CP patients attain MMR and potentially DMR and TFR.

Author Contributions

P.L., J.W. and Y.W. conceived and designed the study. Y.W., S.G., X.C., X.H., Q.L. and L.Z. performed the main experiments and analyzed the data. L.H., P.W., K.L., Y.Z. assisted in collecting the patient samples. Y.W. and P.L. wrote this paper. J.W. and X.D. provided advice and revised the paper.

Funding

This work was supported by the National Natural Science Foundation of China (No. 82200205, No. 82200148), the Project of Administration of Traditional Chinese Medicine of Guangdong Province (20242002) and Medical Scientific Research Foundation of Guangdong Province (B2025364).

Ethics Statement

This study was approved by the Ethics Committee of Guangdong Provincial People's Hospital (KY‐Q‐2021‐284‐01).

Consent

Informed consent was obtained from patients.

Conflicts of Interest

The authors declare no conflicts of interest.

Supporting information

Table S1: Normality testing results.

Figure S1: Representative amplification curves.

Figure S2: Flow cytometry gating strategy.

(A) Lymphocytes were first gated based on forward scatter (FSC) and side scatter (SSC).

(B) CD4⁺ T cells were identified from the lymphocyte population.

(C) The expression of PD‐1 on CD4⁺ T cells was gated on CD4⁺ T cells.

(D) Among CD4⁺ T cells, CXCR5⁺PD‐1⁺ double‐positive cells were defined as T follicular helper (Tfh) cells.

Data Availability Statement

All data generated or analyzed during this study are in the submitted article.