Current serological antibody tests for severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2) require enzyme or fluorescent labels, and the titer well plates cannot be reused. By immobilizing histidine (His)-tagged SARS-CoV-2 spike (S1) protein onto tris‒nitrilotriacetic acid (tris-NTA) sensor and using the early association phase for mass-transfer-controlled concentration determination, we developed a rapid and regenerable surface plasmon resonance (SPR) method for quantifying anti-SARS-CoV-2 antibody. On a five-channel SPR instrument and with optimized S1 protein immobilization density, each of the four analytical channels is sequentially used for multiple measurements, and all four channels can be simultaneously regenerated once they have reached a threshold value. Coupled with a programmable autosampler, each sensor can be regenerated at least 20 times, enabling uninterrupted assays of more than 800 serum samples. The accuracy and speed of our method compare well with those of the enzyme-linked immunosorbent assay (ELISA), and the detection limit (0.057 μg mL) can easily meet the requirement for screening low antibody levels such as those in convalescent patients. In addition, our method exhibits excellent channel-to-channel (RSD = 1.9%) and sensor-to-sensor (RSD = 2.1%) reproducibility. Obviation of an enzyme label drastically reduced the assay cost, rending our method (<60 cents) much more cost effective than those of commercial ELISA kits ($4.4-11.4). Therefore, our method offers a cost-effective and high-throughput alternative to the existing methods for serological measurements of anti-SARS-CoV-2 antibody levels, holding great promise for rapid screening of clinical samples without elaborate sample pretreatments and special reagents.